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Evolution of complex organelles in dinoflagellates Gavelis, Gregory S. 2015

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  EVOLUTION OF COMPLEX ORGANELLES IN DINOFLAGELLATES  by  Gregory S. Gavelis   A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF   DOCTOR OF PHILOSOPHY   in   The Faculty of Graduate and Postdoctoral Studies  (Zoology)          THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)   December 2015   © Gregory S. Gavelis, 2015   ii	Abstract  Dinoflagellates are an abundant and diverse group of aquatic eukaryotes, with members that have photosynthetic, heterotrophic, or mixotrophic life strategies, as well as a number of unique cytological features. My thesis focuses on two groups of closely related dinoflagellates: polykrikoids and warnowiids.  Both include heterotrophic as well as plastid-bearing members, though the number of times photosynthesis has been lost (or gained) in each group is unclear, and the presence and provenance of plastids in some species (e.g., Nematodinium sp. and Polykrikos lebouriae) have been debated. Polykrikoids and warnowiids also contain some of the most complex subcellular structures described--such as nematocysts and, in warnowiids, eye-like ocelloids. Yet these groups are rare in nature and uncultivated, and as such, the origins of their complex organelles are unclear. For my thesis, I modified existing techniques for use on single-cell environmental isolates, and applied these techniques to wild polykrikoid and warnowiid cells. By exploiting the common splice leader sequence found on dinoflagellate transcripts, I was able to amplify a single-cell transcriptome from Polykrikos lebouriae—a dinoflagellate with aberrant plastids. Coupled with single-cell genomics using multiple displacement amplification (MDA), I demonstrated that Polykrikos lebouriae has retained peridinin-type plastids, while photosynthesis has been lost in multiple other polykrikoid species independently.  Using MDA and single-cell transmission electron microscopy, I also determined that the eye-like ocelloid of Nematodinium sp. is made in part from a peridinin plastid, and also from mitochondria. Specifically, single-cell focused ion beam scanning electron microscopy (FIB-SEM) allowed me to demonstrate that a retina-like portion of the ocelloid is a small part of a much larger peridinin-plastid that ramifies throughout the Nematodinium cell.  Lastly, I investigated the evolution of nematocysts in Polykrikos spp. and Nematodinium sp. using a combination of transcriptomics, TEM, SEM, and FIB-SEM, and inferred that “nematocysts” in these groups evolved independently from those in cnidarians.  Thus, nematocyst-like extrusive organelles appear to have evolved multiple times in eukaryotes. The data presented in this thesis show how extreme subcellular  iii	complexity has evolved in dinoflagellates through both endosymbiotic and autogenous origins.     iv	 Preface  Every figure presented in this thesis is my original work, except for the micrographs in Figures 3.1a,b, 3.5, and 3.6b, which were provided by Shiho Hayakawa—the second author of a paper on which Chapter 3 is based. All of these data have been either published or submitted for publication, as described below:  Chapter 2 is based on the following published article: Gavelis, G.S., White, R.A., Suttle, C.A., Keeling, P.J., and Leander, B.S. (2015). Single-cell transcriptomics using spliced leader PCR: Evidence for multiple losses of photosynthesis in polykrikoid dinoflagellates. BMC Genomics. 16:528 (9 pages).  I designed the experiments in collaboration with Dr. Rick White, Dr. Patrick Keeling, and Dr. Brian Leander. I collected and imaged cells, carried out phylogenetic analysis, predicted plastid-targeting peptides, and composed the figures. Together Dr. White and I adapted SLPCR for single cells, carried out the SLPCR and MDA reactions, assembled genomic reads and analyzed the transcriptomic data. I wrote the manuscript with Dr. Leander and all authors subsequently participated in the drafting process.   Chapter 3 is based on the following published article: Gavelis, G.S., Hayakawa, S., White, R.A., Gojobori, T., Suttle, C.A., Keeling, P.J., and Leander, B.S. Eye-like ocelloids are built from different endosymbiotically acquired components. Nature. 523:204-207.  I performed dissected-organelle and single-cell genomics, and phylogenetic analyses on specimens I collected in Canada. Dr. Shiho Hayakawa performed TEM and transcriptomics on specimens she collected in Japan. Dr. Rick White  v	prepared genomic libraries for sequencing and participated in single-cell genomics. I composed the figures and wrote the manuscript with Dr. Brian Leander and Dr. Patrick Keeling.  Chapter 4 is based on the following material currently being drafted for publication.  Gavelis, G.S., Wakeman, K.C., Ripken, C., Herranz, M., Holstein, T., Ozbek, S., Keeling, P.J., and Leander, B.S. Comparative ultrastructure and genomics suggest that nematocysts evolved at least twice. (In preparation).  I conducted the fieldwork, light microscopy, TEM, 3-D reconstructions from FIB-SEM, and data analysis. Dr. Kevin Wakeman prepared single cells for TEM and FIB-SEM, and worked with Christina Ripken on FIB-SEM imaging. Dr. Thomas Holstein and Dr. Suat Ozbek contributed proteomic data from cnidarians. I wrote the preliminary draft of this manuscript, and Dr. Keeling and Dr. Leander provided funding, infrastructure and guidance throughout the project.    vi	Table of contents  Abstract .................................................................................................................. ii Preface .................................................................................................................. iv Table of contents ................................................................................................... vi List of tables .......................................................................................................... ix List of figures ......................................................................................................... x List of abbreviations .............................................................................................. xi Glossary of terms ................................................................................................. xii Acknowledgements ............................................................................................. xiv 1 Introduction ......................................................................................................... 1         1.1 Historical perspective .............................................................................. 1         1.2 The scale and spectrum of eukaryotic complexity .................................. 2         1.3 What are dinoflagellates? ........................................................................ 4         1.4 A circuitous history of plastid loss and acquisition .................................. 5         1.5 The significance of polykrikoids and warnowiids ..................................... 6         1.6 Utility and challenges of genomics and transcriptomics on rare and I       I       uncultivated single cells ................................................................................. 7         1.7 Importance of single-cell microscopy ...................................................... 8         1.8 Approaches and goals ............................................................................. 9 2 Multiple losses of photosynthesis in polykrikoid dinoflagellates are revealed by single-cell transcriptomes .................................................................................... 10         2.1 Introduction ............................................................................................ 10         2.2 Materials and methods .......................................................................... 12                 2.2.1 Collection of organisms .............................................................. 12                 2.2.2 Single-cell transcriptomics .......................................................... 13                 2.2.3 Single-cell genomics .................................................................. 14                 2.2.4 Sequencing, assembly, and annotation ..................................... 14                 2.2.5 Multiple sequence alignments .................................................... 15                 2.2.6 Molecular phylogenetic analyses ............................................... 16                 2.2.7 Presequence analysis of plastid targeted genes ........................ 16  vii	        2.3 Results .................................................................................................. 17                 2.3.1 Genes for plastid-targeted proteins obtained from a single-cell I       I       I       transcriptome ....................................................................................... 18                 2.3.2 Plastid-encoded genes obtained from single-cell genomic data. 18         2.4 Discussion ............................................................................................. 23                 2.4.1 Utility of transcriptomics and genomics on single cells .............. 23                 2.4.2 Peridinin plastids in Polykrikos lebouriae ................................... 24                 2.4.2 Hypothesis for polykrikoid plastid evolution ............................... 24                 2.4.2 Future applications of SLPCR .................................................... 25 3 Eye-like ocelloids in warnowiid dinoflagellates evolved from different endosymbiotically acquired components ............................................................ 28         3.1 Introduction ............................................................................................ 28         3.2 Material and methods ............................................................................ 29                 3.2.1 Collection of organisms .............................................................. 30                 3.2.2 Flourescence and DIC microscopy ............................................ 30                 3.2.3 Single-cell TEM of Nematodinium sp. and Erythropsidinium. sp.30                 3.2.4 Transcriptomics on Warnowia sp. and Erythropsidinium sp. ..... 31                 3.2.5 Isolation of the retinal bodies of Nematodinium sp.. .................. 31                 3.2.6 Single-organelle genomics of Nematodinium sp. ....................... 31                 3.2.7 Molecular phylogenetic analyses ............................................... 32                 3.2.8 Focused ion beam scanning electron microscopy (FIB-SEM) ... 33         3.3 Results .................................................................................................. 34                 3.3.1 TEM in Nematodinium sp. and Erythropsidinium sp .................. 34                 3.3.2 Transcriptomics in Erythropsidinium sp. and Warnowia sp. ....... 38                 3.3.3 Single-organelle genomics in Nematodinium sp. ....................... 39                 3.3.4 3-D reconstruction of the ocelloid in Nematodinium sp. ............. 41         3.4 Discussion ............................................................................................. 46 4 Comparative ultrastructure and genomics suggest that nematocysts evolved at least twice ............................................................................................................ 48         4.1 Introduction ............................................................................................ 48         4.2 Materials and methods .......................................................................... 49  viii	                4.2.1 Genome and transcriptome mining for nematogenic proteins ... 49                 4.2.2 Collection of organisms .............................................................. 50                 4.2.3 Inducing nematocyst discharge .................................................. 50                 4.2.4 Scanning electron microscopy on isolated organelles ............... 51                 4.2.5 Single-cell transmission electron microscopy ............................. 51         4.3 Results .................................................................................................. 51                 4.3.1 No predicted dinoflagellate proteins with affinity to cnidarian I       I       I       nematocyst machinery ........................................................................ 51                 4.3.2 SEM and TEM illustrate a lack of homology between polykrikoid I       I       I       and cnidarian nematocysts .................................................................. 55                 4.3.3 TEM and FIB-SEM reveals new levels of complexity in the I       I   I       I       nematocysts of Nematodinium sp. ...................................................... 58         4.4 Discussion ............................................................................................. 63                 4.4.1 Independent origins of pressurized ballistics ............................. 63                 4.4.2 Complexity through duplication .................................................. 64                 4.4.3 Concluding remarks ................................................................... 64 5 Conclusion ........................................................................................................ 67         5.1 Contributions to understanding the evolution of complex dinoflagellate          I       organelles .................................................................................................... 67         5.2 Future directions .................................................................................... 70 Bibliography ......................................................................................................... 73  		 	 ix	  List of tables  Table 2.1 Nuclear-encoded plastid-targeted genes transcripts ........................... 21    x	List of figures  Figure 2.1 Phylogeny of polykrikoid dinoflagellates ............................................ 10 Figure 2.2 Basic steps in splice leader primed PCR (SLPCR) ............................ 15 Figure 2.3 Transcripts expressed by a single-cell of Polykrikos lebouriae .......... 18 Figure 2.4 Phylogenetic affinity of plastid genes in Polykrikos lebouriae ............ 19 Figure 2.5 Peridinin type plastid-targeting sequences ........................................ 20 Figure 3.1 Light micrographs (LM) of warnowiids used in this study .................. 25 Figure 3.2 Ultrastructure of the retinal body in Nematodinium sp. ...................... 31 Figure 3.3 Development in warnowiids ............................................................... 32 Figure 3.4 Thylakoid forms in Nematodinium sp. ................................................ 33 Figure 3.5 Transient thylakoids in the retinal body as viewed with transmission electron microscopy ............................................................................................ 34 Figure 3.6 Genomics and structure of organelles in the ocelloid ........................ 36 Figure 3.7 Phylogeny of retinal-body encoded proteins. ..................................... 37 Figure 3.8 Three-dimensional reconstruction of the ocelloid of Nematodinium sp. using FIB-SEM tomography. ............................................................................... 39 Figure 3.9 FIB-SEM slices of the cornea-like layer of mitochondria in the ocelloid of Nematodinium sp. ........................................................................................... 40 Figure 3.10 Inferences on the predatory nature of Nematodinium sp. from live and preserved specimens. .................................................................................. 41 Figure 4.1 Nematocysts in Polykrikos kofoidii and Nematodinium sp. ................ 45 Figure 4.2 Distribution of extrusomes in eukaryotes ........................................... 46 Figure 4.3 Distribution of cnidocyst machinery in eukaryotes ............................. 51 Figure 4.4 Nematocysts in Polykrikos kofoidii ..................................................... 53 Figure 4.5 Nematocysts in Nematodinium sp. .................................................... 54 Figure 4.6 Comparison of nematocyst characteristics in cnidarians and dinoflagellates ..................................................................................................... 55 Figure 5.1 Dinoflagellate eyespot types within a phylogenetic context ............... 63 	  xi	List of abbreviations    AtpA – Plastid gene encoding synthase CF1 alpha chain protein.  CAMERA – Community cyberinfrastructure for advanced microbial ecology research and analysis.  LGT – Lateral gene transfer.  LSU – Large (28s) subunit ribosomal rDNA gene.  ML – Maximum likelihood.  MDA – Multiple displacement amplificiation.  PetB – Plastid gene encoding cytochrome B6 protein.  PetD – Plastid gene encoding cytochrome B6/F complex subunit 4 protein.  PsaA – Plastid gene encoding photosystem I P700 chlorophyll a apoprotein A1.  PsaB – Plastid gene encoding photosystem I P700 apoprotein A2.  PsbA – Plastid gene encoding photosystem II protein A1.  PsbB – Plastid gene encoding photosystem II CP47 protein.  PsbC – Plastid gene encoding photosystem II CP43 protein.  PsbD – Plastid gene encoding photosystem II protein D1  SL – Spliced leader.   SLPCR – Spliced leader based polymerase chain reaction.  SSU – Small (18s) subunit ribosomal rDNA gene.          xii	Glossary of terms    Docidosome – A putative ballistic organelle type found in the dinoflagellate Actiniscus pentasterias. Each docidosome has a bilateral symmetry that suggests a “double-barreled” discharge mechanism.  Extrusomes – Ballistic secretory organelles. Many protists use extrusomes for predation or defense.   Helicosporidians – A highly-derived parasitic group of green algal heritage.  To initiate infection, a helicosporidian cyst rapidly unrolls a filamentous cell that penetrates the host epidermis.   Kleptoplastids – Plastids that have been taken from a prey organism and function temporarily within the predator.   Microsporidians – Unicellular parasites derived from fungi. Microsporidians use a “polar filament” to inject their cytoplasm into host cells with explosive force.  Minicollagens – A family of cnidarian proteins that share a collagen-triple helix domain, as well as N and C terminal cystein rich domains that allow adjacent minicollagens to crosslink thorugh disulphide bridges. Minicollegens form up to 70% of cnidarian nematocysts by dry weight, comprising most of the capsule and tubule.  Mixotrophy – A feeding strategy that combines both autotrophy and heterotrophy. Most photosynthetic dinoflagellates are also mixotrophic.  Myxozoans – Unicellular cnidarians that have evolve a parasitic lifestyle.  Myxozoans use “polar filaments” (homologous to nematocysts) to attach to their host prior to invasion.  Mucocysts – Ballistic organelles found in some ciliates and dinoflagellates (though not necessarily monophyletic).  Mucocysts can discharge a mucilaginous material which surrounds the cell as a defensive hyaline cyst.  Ocelloid – An eye-like structure found in warnowiid dinoflagellates. Though the ocelloid possesses structures resembling a lens, iris-like rings, and a pigmented retinal body, it is subcellular, and therefore consists of organelles.  Operculum – A hatchlike structure that covers the opening of nematocysts in cnidarians.  Peridinin plastids – Red algal derived plastids that are an ancestral feature for  xiii	dinoflagellates. They share a common origin with the plastids in chromerids and apicoplasts in apicomplexans.  PhiX – A bacteriophage. When preparing DNA libraries for sequencing, PhiX DNA is often used as a control sample from which the unknown amount sof DNA in other samples can be estimated.  Pseudocolony – A unique cytological arrangement found in cells of the dinoflagellate genus Polykrikos.  Each pseudocolony is a cell with two nuclei, and between four and eight segments (or zooids) that resemble partially-divided cells arrested at cytokinesis. Each pseudocolony is motile, has from 8 to 16 flagella, and otherwise behaves like a typical dinoflagellate cell.  Retinal body – A cup-shaped black/red pigmented body in the ocelloid of warnowiids. Our findings suggest that it is part of a highly modified peridinin plastid.  Stylet – Piercing structures in the nematocysts of cnidarians. Generally arranged in pairs, stylets puncture the cuticle or epidermis of the target organism, and aid in the delivery of toxins.  Taeniocysts – Toxicyst-like organelles found in most Polykrikoids. They are arranged in a series with the nematocysts, in the forwardmost position.  Toxicysts – Extrusive organelles that deploy a syringe-like tube from within a capsule.  As their name suggests, toxicysts seem to serve as rapid toxin delivery systems, and are generally employed in predation.  Trichocysts – Seemingly defensive ballistic organelles found in some ciliates, colpodellids, and most dinoflagellates. Discharge occurs when these paracrystalline rods rapidly repolymerize into a longer ribbonlike form.   Warnowiids – Unarmored dinoflagellates characterized by eyelike structures called ocelloids (see ‘Ocelloid’).  Zooid – See ‘Pseudocolony’            xiv	Acknowledgements  This work was made possible by a four-year fellowship from the Department of Zoology, as well as support and guidance from by Dr. Brian Leander and Dr. Patrick Keeling. The technical knowledge of Dr. Gethin Owen, Derrick Horne, Erick James, and Garnet Martens was also instrumental to the project.  I thank Dr. Kevin Wakeman for introducing me to single cell TEM, and for patiently tolerating me even after I launched his new truck off a precipice and into the high intertidal zone at Clover Point, Victoria.  I dedicate this thesis to my wife, who is an amazing field assistant and has kept us both alive.   1	1 Introduction  1.1 Historical Perspective  Historically, there has been resistance to the idea that complexity can arise from, or manifest within, a single cell. Early biologists could not fathom how a cell as “simple” as a sperm or egg contained the instructions to build a human being. Some found it easier to believe that miniature, nascent humans, or “homunculi,” were tucked inside spermatozoa (recounted in Beaty 1985); indeed, several naturalists claimed to have seen homunculi under the light microscope! Eventually this illusion was dispelled by skepticism and better optics, and scientists were forced to reckon with the fact that animals do indeed develop from single cells. But the processes governing development were largely mysterious, as were the inner workings of the cell.  A century before the discovery of genes and DNA, Oskar Hertwig determined that the nucleus is the vehicle of (most of) our heritable traits. Prior to his observations, it was unclear which, if any, specific part of the cell contained our genetic legacy. While studying external fertilization in sea urchins, Hertwig noticed that the sperm pronucleus fused with the pronucleus of the egg, and thus presented the first compelling mechanism for how heritable features are transmitted between generations.   In 1884, during one trip to the Mediterranean to collect sea urchin gametes, Hertwig encountered a cell more peculiar than any yet described. He had sifted the water column with a plankton net, hoping to gather sea urchin sperm and eggs, and then observed the filtrate under the light microscope. Among the plankton and spermatozoa was a larger flagellated cell, one that bounced up and down on a piston-like appendage. Stranger still, the cell seemed to behold him with an eyelike structure bearing a lens, iris, and pigmented retina. Finding it remarkable that a single cell could have an “eye,” Hertwig tried unsuccessfully to  2	fix the specimen in glutaraldehyde, and ultimately published several detailed illustrations that he had drawn from memory (Hertwig 1884). Hertwig’s findings were swiftly criticized by the zoologist Karl Vogt. Vogt argued that a cell cannot possess an eye, given that eyes are made of cells. The most likely explanation, Vogt stated, was that Hertwig had described a horribly contorted ciliate in its death throes—one that had eaten the eye of a jellyfish. While he responded to several rounds of Vogt’s acrimonious rebuttals, Hertwig never encountered another specimen, and his strange cell was considered by many to be apocryphal (described in Kofoid and Swezy 1920).   The specimen that Hertwig described is now known to belong to a protist group called the dinoflagellates—specifically the warnowiids—and its eyelike ocelloid is still the subject of speculation. In the 1970s, Claude Greuet applied transmission electron microscopy to warnowiids, and found that the pigmented component of the ocelloid (the “retinal body”) contained membranes reminiscent of a plastid (Greuet 1971, 1977, 1987). Therefore, Greuet hypothesized that ocelloid developed, at least in part, from a highly modified plastid. More recently, Walter Gehring, known for his lab’s discovery of the Hox genes that are developmentally critical in animals, hypothesized that the ocelloid’s eyelike nature is not incidental, but is homologous to the eyes of animals (Gehring 2004).   This debate could be settled by a molecular analysis of ocelloid machinery, but prior to this thesis, such work was largely prevented by the rarity and fragility of warnowiid cells. The evolutionary origin of the ocelloid is one of several mysteries that I addressed in the course of my doctoral research, and document here, as an effort to understand the evolution of complexity in this bizarre group of eukaryotic organisms—the dinoflagellates.      3	1.2 The scale and spectrum of eukaryotic complexity  There is a spectacular range of complexity across eukaryotic cells, from parasites that have lost their mitochondria and thousands of nuclear genes (Corradi et al. 2007), to highly differentiated cells, such as ciliates, radiolarians, and multinucleated organisms that challenge the definition of unicellularity (Leander, Saldarriaga and Keeling 2002). Yet all eukaryotes—including plants, animals, fungi, seaweeds and protists—share a common ancestor that had a nucleus, endomembrane system, cytoskeleton and mitochondria. These features differentiate eukaryotes from bacteria and archaea, which have no nuclei and few if any other compartments within the cell. How did eukaryotes become so complex?    Phylogenomic comparisons suggest that several eukaryotic characteristics arose “autogenously” through the duplication and differentiation of pre-existing features, such as the divergence of the nuclear envelope, endoplasmic reticulum, and Golgi bodies from a single ancestral compartment (Dacks and Field 2007), or—as a molecular example—the invention of the cytoskeleton using proteins inherited from prokaryotic ancestors, which initially had non-cytoskeletal functions (such as the FtsZ protein used for septum formation in bacterial division, which is homologous to eukaryotic tubulins: Koonin 2015).   The second major route in cellular evolution is an “exogenous” one—i.e., involving the acquisition of components from foreign lineages. The most famous examples are mitochondria and plastids, which were once bacteria that became harnessed during endosymbiosis (Keeling 2010, Mast et al. 2014)—and plastids have since been transferred even farther afield through additional rounds of endosymbiosis (Archibald 2012, Yamaguchi, Yubuki, and Leander 2012). Genes too can have exogenous origins via lateral gene transfer. By definition, lateral gene transfer occurs between distantly related organisms and can also occur between different genomes in the same cell (e.g., nuclear, plastid, and  4	mitochondrial genomes) (Koonin 2015). Even viruses have been a source of biological novelty in some eukaryotes, such as the viral nucleoprotein that seems to be involved in DNA packing within the nucleus of dinoflagellates (Gornik et al 2012). Features that add biological complexity may not initially be adaptive (or necessarily ever become efficient or useful; Keeling, Leander, Lukeš 2010), and causal factors for the vast range in complexity across eukaryotes remain unclear.   With a few exceptions (e.g., ciliates), cell biology has focused on relatively simple—and therefore tractable—model organisms, because the streamlined biology of durable, rapidly-multiplying organisms such as yeast or E. coli is more accessible than the giant genomes of amoebae or the dizzying architecture of radiolarians. As such, our knowledge of rare and uncultivated groups lags far behind these models, and the upper limits of cellular complexity remain poorly explored. This is true even for ecologically important organisms such as dinoflagellates, which are both complex and abundant—being the most abundant eukaryotic phytoplankton after diatoms (though certain dinoflagellate taxa are quite rare).  1.3 What are dinoflagellates?  Dinoflagellates can be recognized by (and were named after) their helical swimming pattern, as each dinoflagellate is propelled by an undulating “transverse” flagellum that girdles the cell, and steered by a “longitudinal” flagellum that trails behind it. This flagellar arrangement—along with a large birefringent nucleus (“dinokaryon”)—are the diagnostic features of the Dinophyceae (Kofoid and Swezy 1921, Taylor 1987). Remarkably, the dinokaryon contains permanently condensed chromosomes, in which heterochromatin lacks histones and is packed so densely as to approach a “liquid crystalline” state (Liu and Kattawar 2013). This bizarre arrangement inspired early taxonomists to split dinoflagellates into their own domain of life: the Mesokaryota, which was thought to have emerged before the eukaryotic  5	invention of histones (reviewed in Taylor 1987). Since then, molecular phylogenetic studies have firmly placed dinoflagellates as eukaryotes within the Alveolata, as sisters to a clade comprised of apicomplexans (parasites such as Toxoplasma) and chrompodellids (photosynthetic coral reef endosymbionts and some free-living predators) (Janouskovec et al. 2015). In other words, dinoflagellates do not represent a “missing link” in cellular evolution, but are a more recent experiment in eukaryotic organization.  The cytological strangeness of dinoflagellates is not isolated to the nucleus. As we will see, it reaches across several organelle systems.  1.4 A circuitous history of plastid loss and acquisition  Dinoflagellates are among the most common eukaryotic phytoplankton; slightly less abundant than diatoms, and more prevalent than haptophytes. Plastids in these groups generally have chlorophylls a and c, and were introduced through “secondary endosymbiosis” (Keeling 2010)—that is, the ancestor(s) of diatoms, dinoflagellates, and haptophytes acquired photosynthesis by ingesting red algal cells. While it is unclear whether secondary symbiosis occurred once or independently for each of these groups, the red algal cells were ultimately dispensed with, leaving behind plastids that integrated with their new hosts. In dinoflagellates, these “peridinin” plastids have transferred the vast majority of their genes to the nucleus, and the remaining genes (located on plasmid-like minicircles) constitute the smallest plastid genomes of any known photosynthetic organisms (Howe, Nisbet and Barbrook 2008). Stranger still, several dinoflagellate lineages replaced their peridinin plastids with those from green algae (through “serial secondary symbiosis”) (Minge et al. 2010), or from other diatoms, haptophytes, and cryptophytes (through “tertiary symbiosis”) (Imanian, Pombert, and Keeling 2010, Archibald 2012).   The loss of photosynthesis is prevalent in dinoflagellate evolution, as roughly half of all species have become heterotrophic (Saldarriaga et al. 2001). The rest are  6	mostly mixotrophic species (utilizing both phototrophy and heterotrophy)—as they generally supplement photosynthesis with predation.  Elaborate predatory adaptions are especially conspicuous in polykrikoid dinoflagellates (Hoppenrath and Leander 2007b, Matsuoka 2000).  1.5 The significance of polykrikoids and warnowiids  Along with noctilucoids, polykrikoid and warnowiid dinoflagellates have the special distinction of having been initially confused as animals, due to their highly derived forms.  Polykrikoids were initially classified as polychaete larvae, based on their large size and superficially segmented appearance (Kofoid and Swezy 1921 and citations therein). These segments actually represent incompletely divided cells (or zooids), which do not pinch apart but remain fused as a single multinucleate cell or “pseudocolony.”  These large, peculiar cells also possess up to 16 flagella, and probably evolved through one or more rounds of arrested cell division.   Warnowiids are closely related to polykrikoids, as they both branch within the Gymnodiniales sensu stricto (Hoppenrath and Leander 2007a,b, Hoppenrath et al. 2009). While they lack pseudocolonies, warnowiids possess a peculiar defining feature—the ocelloid.  This eye-like structure (described in section 1.1) bears features reminiscent of a lens, irises and a pigmented retinal body, and was initially suspected to be an artifact produced when the cell scavenged the eye from a jellyfish (recounted in Kofoid and Swezy 1921). However, TEM studies have firmly established the ocelloid as a distinct multi-organellar complex (Greuet 1977). There is some ultrastructural evidence that its pigmented retinal body is a highly modified plastid (Greuet 1989), but this hypothesis is unconfirmed, and at present, the evolutionary origins of the different parts of the ocelloid are unclear.    Due to their extreme rarity and fragility (which generally prevents live observation), virtually nothing of warnowiid life history is known. They are  7	presumed to be heterotrophs, based on the absence of plastids in most species (excluding Nematodinium sp. in which the presence of plastids has been contested; Mornin and Francis 1967, Hoppenrath et al. 2009]). The history of plastid evolution is also unclear in polykrikoids, as roughly half of them lack phototrophy, and those with plastids either have peridinin-type plastids typical of dinoflagellates (Qiu et al. 2013) or have morphologically aberrant plastids—as in Polykrikos lebouriae—in which the membranous ultrastructure is more reminiscent of plastids from diatoms or haptophytes (Hoppenrath and Leander 2007a). Many groups are probably mixotrophic, such as those with both plastids and predatory organelles: the nematocysts.    Nematocysts are incredibly complex predatory extrusomes that are found in most polykrikoids, and also in the warnowiid Nematodinium—albeit with a very different ultrastructure (Greuet 1971). As with plastids in these groups, the homology between these different nematocysts types remains unclear. In addition to nematocysts, polykrikoids possess a unique organelle type; the taeniocyst, which resembles a very elaborate sort of toxicyst, thought its function is unknown.  In polykrikoids, the taeniocyst is always arranged in the cell with a nematocyst close behind it. Whether these two extrusomes work in synchrony during prey capture is entirely unclear. This constellation of complex traits makes warnowiids and polykrikoids fascinating groups in which to study character evolution and explore the mechanisms that produce subcellular complexity in eukaryotes.  1.6 The utility and challenges of genomics and transcriptomics on rare and uncultivated single cells  Our limited understanding of polykrikoids, warnowiids, and other rare protists reflects a dearth of tools available for studying uncultivated cells. Approaches using PCR—the standard method for genetic analysis of individual cells—require background knowledge in order to design primers for the desired gene  8	sequences, and there is little groundwork for this in most dinoflagellate groups. Moreover, it is especially challenging to design plastid PCR primers for dinoflagellates, given the rapid sequence evolution of these plastid genes (Bachvaroff, Sanchez-Puerta, and Delwiche 2006). Non-specific genomic amplification offers an alternative to PCR, as employed in multiple displacement amplification (MDA), which uses random primers. Significant advances in single cell genomics have been made using MDA, but due to its nonspecificity, MDA will also amplify undesired contaminant sequences (Yoon et al. 2011). For this reason, MDA is not ideal for investigating novel symbioses or plastid acquisition, as it is difficult to determine if the amplified reads originated from the symbiont or from genetic contaminants introduced from prey cells or the environment. Transcriptomics offers an alternative (or complementary) approach to genomics—especially among dinoflagellates in which the genome sizes are often prohibitively large (Shoguchi et al. 2013). Yet, like MDA, total-RNA sequencing is apt to capture contaminant sequences that can confound evolutionary analyses. In the next two chapters, I describe our experiments with new permutations of existing genetic tools, including transcriptomics and genomics, and explain my findings from single uncultivated cells, and even individual organelles.  1.7 Importance of single-cell microscopy.  Some of the most surprising findings in evolutionary biology have recently come from genomic approaches, such as environmental sequencing surveys (metagenomics) or from single-cell isolates. Landmark genomic studies have uncovered previously overlooked lineages and added large branches to the tree of life at some evolutionarily important junctures, such as between parasites and free-living algae (Janouskovec et al. 2013,) or at the nexus of eukaryotes and archaea (Spang et al. 2015). But even from a complete genome, we cannot accurately predict a microorganism’s external appearance, internal anatomy, or behavior.    9	Major errors and misinterpretations are also more likely to be overlooked in solely sequence-based studies, for lack of other approaches to confirm the findings. Indeed, flawed analyses resulting from sequence contamination have caused some organisms to be phylogenetically misplaced, such as the erroneous placement of the new phylum Xenoturbellida among the mollusks, when it is now known to be of a deuterosome affinity (Bourlat et al. 2006)—due to genetic contamination from bivalve prey, or the false designation of “picobilophytes” as phytoplankton—evidently due to genetic contamination from cyanobacteria—given that more recent whole-genome reconstructions show them to be nonphotosynthetic (Yoon et al. 2011). Therefore, it is important to complement sequence-based approaches with other methods—such as microscopy—in order to validate the findings and also place them within a clear cellular context.  In Chapters 3 and 4 of this thesis, I experimented with advanced microscopic techniques such as TEM and FIB-SEM, and show here that they are applicable to rare single cells.   1.8 Approaches and goals  My thesis research explores the biology of rare and uncultivated dinoflagellates, particularly those with complex organelles. Here I use a combination of both established and novel applications of transcriptomics, genomics, and electron microscopy on individual cells or organelles, with the goal of understanding (1) the pattern of plastid evolution in polykrikoid dinoflagellates, (2) the evolutionary origin of the warnowiid ocelloid, and (3) the potential homology or analogy between extrusive “nematocysts” in warnowiids, polykrikoids, and cnidarians.    10	2 Multiple losses of photosynthesis in polykrikoid dinoflagellates are revealed by single-cell transcriptomes.  2.1 Introduction  Most microbial eukaryotes are uncultivated and thus poorly suited to standard genomic techniques. This is the case for Polykrikos lebouriae, a dinoflagellate with ultrastructurally aberrant plastids. It has been suggested that these plastids stem from a novel symbiosis with either a diatom or haptophyte (Hoppenrath and Leander 2007a) but this hypothesis has been difficult to test as P. lebouriae dwells in marine sand rife with potential genetic contaminants.  Early-branching polykrikoids, such as Polykrikos geminatum and P. hartmanii, have plastids with three membranes and triple-stacked thylakoids that are characteristic of the secondary peridinin-type plastids of most dinoflagellates (Qiu et al. 2013, Hoppenrath et al. 2010) (Fig. 2.1). The plastids of P. lebouriae, however, are reportedly enveloped by only two membranes (a trait that is more consistent with primary plastids), and to contain double-stacked thylakoids similar to those found in haptophytes and diatoms (Hoppenrath and Leander 2007a). P. lebouriae also has a conspicuous phylogenetic position, as a plastid-bearing mixotroph nested among three heterotrophic species (P. herdmanae, P. schwartzii and P. kofoidii) (Fig. 2.1) (Hoppenrath and Leander 2007b).  Leander and Hoppenrath (2007b) interpreted this as evidence of either multiple losses of photosynthesis among P. herdmanae, P. schwartzii and P. kofoidii, or a single loss at the base of this group, followed by acquisition of tertiary plastids in P. lebouriae from a diatom or haptophyte. These hypotheses remain untested, as several attempts to cultivate P. lebouriae have been unsuccessful (Aika Yamaguchi, Mona Hoppenrath, personal communication), and PCR amplification of plastid genes in P. lebouriae has consistently failed with PCR primers used successfully in other taxa.   11	 Multiple displacement amplification (MDA) is a powerful tool for whole-genome amplification from small amounts of template DNA, but it is nonspecific and therefore prone to contamination.  We employed this technique to amplify a partial plastid genome of Polykrikos lebouriae, and supplemented this with a dinoflagellate-specific transcriptomic approach, both to ensure that our plastid amplification did not stem from non-dinoflagellate environmental contaminants (i.e.; diatoms, haptophytes or other algae that share the same habitat as P. lebouriae), and to test whether the plastids are functionally integrated into the cell (ie; if the nucleus expresses plastid-targeted genes) rather than being simply retained as kleptoplastids. We synthesized cDNA from single P. lebouriae cells, which we primed for PCR with a 21 bp spliced leader sequence specific to dinoflagellates, via SLPCR (Fig. 2.2).  Previous researchers have established the effectiveness of SLPCR for amplifying dinoflagellate transcripts from a large volume of wild-caught plankton or coral tissue (Zhang et al. 2007, 2013), and this is the first study to apply SLPCR at the scale of single cells.     MDA and SLPCR allowed us to illuminate regions of the plastid genome in P. lebouriae as well as nuclear gene expression.  In concert, these methods provided evidence of the presence and provenance of functional plastids in P. lebouriae, and allowed us to test hypotheses for plastid evolution in this uncultivated group.  12	 Figure 2.1. Phylogeny of polykrikoid dinoflagellates. Maximum likelihood (ML) tree inferred from the 31-taxon alignment (1,915 unambiguously aligned sites) of concatenated small and large ribosomal rDNA sequences using the GTR + Γ substitution model (14 basal taxa not shown due to size constraints, for original tree see Gavelis et al. 2015b).  Bootstrap support values 65 or higher and Bayesian posterior probabilities are listed above each branch. The illustrations depict the pseudocolonies of polykrikoid species; orange indicates a photosynthetic pseudocolony, and gray indicates a non-photosynthetic pseudocolony.      13	  2.2 Materials and methods   2.2.1 Collection of organisms.  After several fruitless searches for Polykrikos lebouriae in its previously reported site in Centennial Beach, BC, broader sampling was conducted on the Oregon coast. In early October, samples of the upper 1-cm of marine sand were collected during low tide from the mid intertidal zone in Cannon Beach, Oregon (45.886346, -123.965043). Polykrikoid-type cells were noticed in these samples using a pocket microscope. Within 36 h, the samples were transported to the University of British Columbia, kept in an open dish of moist sand and exposed to natural day/night rhythms. Uhlig's seawater ice method (Uhlig, Thiel, and Gray 1973) was used to draw cells from the sand into a Petri dish, where they were collected individually by micropipette.  Cells were visually identified based on the presence of two-nuclei, eight zooid segments, and plastids, and later confirmed through analysis of the large subunit (LSU) ribosomal RNA gene. To reduce the chance of genetic contamination from prey, we selected cells of P. lebouriae in which no food vacuoles were evident.  Pseudocolonies of P. lebouriae were washed five times, once in filtered seawater, twice in 1X phosphate buffered saline, and twice in distilled nuclease-free water. For transcriptomics, an individual cell was processed for RNA extraction immediately; five other single-cell isolates were frozen at -80°C. Later that week, the five samples were thawed and pooled for genomics processing using MDA.   2.2.2 Single-cell transcriptomics.  For cell lysis, 0.5 µl of proteinase K was added to the tubes, followed by incubation at 65°C for 10 min, and denaturation at 90°C for 2 min, then rapid cooling at 4°C. First strand cDNA was primed using a GeneRacer OligodT primer  14	(1 µl at 10 µM) RNAse Out (0.5 µl), dNTPs – (2 µl at 10 µM) and incubated at 65 °C for 5 min. After this annealing step, DTT (1 µl at 10 µM), Rnase Out (0.5 µl), and 1 µl of Superscript III reverse transcriptase (Life Technologies, Carlsbad CA) were added and incubated according to the manufacturer’s protocol, allowing for reverse transcription, along with T4 gene 32 (0.5 µl) to maximize contact between the reverse transcriptase and RNA template.  Afterwards, DNA/RNA hybrids were removed with 1 µl of RNAseH, incubated 37 °C for 20 min.  Polyadenylated transcripts were amplified with a a GeneRacer 3’ nested primer (5’-CGCTACGTAACGGCATGACAGTG-3’), and dinoflagellate specificity assured with the a dinoflagellate spliced-leader primer (5′-TCCGTAGCCATTTTGGCTCAAG-3′) and  (Life Technologies, Carlsbad CA).  Thermocycling proceeded through a “touchdown PCR” program, as this was effective for Zhang et al. (2007).  This program progressed through 95°C for 20 s, 72 °C for 2.5 min for 5 cycles; 95 °C for 20 s, 65 °C for 30 s, 72 °C for 2 min for 5 cycles; 95 °C for 20 s, 60 °C for 30 s, 72 °C for 2 min for 5 cycles; and 95 °C for 20 s, 58 °C for 30 s, 72 °C for 2 min for 25 cycles.  Because we were amplifying from a single cell, our PCR reaction program had ten more amplification steps than that of Zhang et al. (2007).  In order to reduce amplification bias, we divided each SLPCR reaction into eight sub-reactions, which ran in parallel, and were pooled at the end.  Reads were quality checked using a Bioanalyzer (Agilent, Santa Clara, CA).     2.2.3 Single-cell genomics.  A frozen cell was thawed on ice then lysed as above.  Multiple Displacement Amplification (MDA) was carried out using the Repli-G mini kit (Qiagen, Venlo, Linburg, Netherlands) as per manufacturer’s instructions, at the maximum recommended time of 15 hours.  The reaction was divided into four sub-reactions to minimize amplification bias and then pooled at the end.  Genomic DNA was amplified non-specifically, including the plastid genes of P. lebouriae.   15	 2.2.4 Sequencing, assembly and annotation.  DNA for libraries was sheared to ~300-400 bp by a Covaris Ultrasonicater (M220) using the manufacturer’s protocol (Covaris, Woburn, MA). Libraries were indexed with TruSeqTm adapter barcodes using Lucigen NxSeq library prep without PCR enrichment to avoid amplification bias.  Libraries were washed with two rounds of AMPureXP magnetic beads (Beckman Coulter, Danvers, MA) at a beads to DNA ratio of 0.8:1 to remove free adapters by size screening.  To ensure sufficient adapter ligation, a sample of the libraries were tested with real-time qPCR (primed to the library indexes), and measured against a digital standard curve [31]. Libraries were screened for purity using a Nanodrop (ThermoFisher, Wilmington, DE) and length and purity using a Bioanalyzer HighSens DNA chip (Agilent, Palo Alto, CA).  Libraries were sequenced with 250 bp paired-end reads on an Illumina MiSeq (GenoSeq UCLA Los Angeles, CA).  A phiX library was used as a quality standard during sequencing.  From the output sequences, phiX was screened and removed, paired ends were merged (if overlapping >30 bp), and non-overlapping reads were interleaved.  Merged reads were checked for a minimum Qscore (Q>30). De novo assembly was performed with Ray (Stamatakis 2006) using a variety of kmer sizes, with 31 chosen as the optimal kmer size for assembling our genomic reads and 53 for reads from SLPCR. Resulting contigs were uploaded to the MG-RAST server, which performed automated annotations and protein predictions (Meyer et al. 2004).  2.2.5 Multiple sequence alignments.  Several alignments were constructed in this study for molecular phylogenetic analysis.  For analysis of ribosomal genes, we concatenated SSU and LSU rDNA sequences, and aligned them across 31 unarmoured dinoflagellates, with Akashiwo sanguinea as the outgroup.  This nucleotide alignment consisted of  16	1,915 unambiguously aligned sites, once gaps and ambiguously aligned regions were removed.  A second alignment was assembled for LSU rDNA sequences alone, in order to confirm that the single cell isolate was Polykrikos lebouriae.  This alignment included 25 dinoflagellate taxa, with 1,229 unambiguously aligned bases.   The remaining alignments were for protein analyses, translated plastid genes PsaA, PsbC and AtpA.  Predicted proteins were aligned with MUSCLE, followed by removal of gaps and ambiguously aligned bases. Using 100 boostraps of RAxML and the substitution model PROTGAMMA, preliminary trees were generated from MUSCLE (Edgar 2004) alignments of 715 amino acids for PsaA, 453 aligned amino acids for PsbC, and 427 aligned amino acids for AtpA.  Having validated these proteins as dinoflagellate plastid-type proteins, we manually concatenated these three alignments into a supermatrix with 1,595 unambiguously amino acids for the final analysis. This alignment incorporated 44 taxa, including representatives of all major groups of photosynthetic eukaryotes, including glaucophytes, red algae, green algae, land plants, haptophytes cryptophytes, stramenopiles, dinoflagellates with peridinin-type plastids, as well Lepidodinium, Karenia, Karlodinium, Durinskia, Kryptoperidinium and six species of cyanobacteria. We chose dinoflagellate taxa for which two or more of the plastid proteins were available in Genbank or CAMERA.  Among dinoflagellates, Symbiodinium, Togula, Lingulodinium, Lepidinodinium, Kryptoperidinium, Durinskia, Aphidinium carterae, Heterocapsa rotundata, and Polykrikos lebouriae had all three proteins, and dinoflagellates with two proteins were incorporated as they led to higher resolution of the dinoflagellate relationships.   2.2.6 Molecular phylogenetic analyses.  Maximum likelihood analysis was run with 1,000 bootstraps using RAxML and PROTGAMMAJTT or GRTGAMMA substitution models for protein and nucleotide sets, respectively (Stamatakis 2001). Bayesian posterior probabilities  17	were calculated for all alignments using the following parameters on the program MrBayes 3.2.2 (Huelsenbeck and Ronquist 2001); (GTR [Lset nst = 6]; gamma distribution [of rate among sites]; and Monte Carlo Markov Chains [starting trees = 4; heating (nchains = 4), default temperature = 0.2; generations = 6,000,000; sample frequency = 100; prior burn-in = 500,000 trees].      2.2.7 Signal peptide analysis of plastid targeted genes.  The N-terminal region of nuclear encoded, putatively plastid targeted genes was analysed for signal peptides with the Hidden Markov Model of SignalP3.0 (Nielsen et al. 1998, Bendtsen 2004) using default settings. Transmembrane helices were predicted using TMHMM v.2.0 (Sonnhammer et al. 1998), and their hydrophobicity scores were calculated with the Kyte-Doolittle amino acid scale from Protscale (http://web.expasy.org/protscale/, last accessed April 22, 2015) using default settings.  Protein sequences were manually aligned in Mega 5.2.2 (Tamura et al. 2013), and imported into Jalview (Clamp 2004), where the charge and hydrophobicity of amino acids were color-coded (Fig. 2.5).    18	 Figure 2.2 Basic steps in spliced leader primed PCR (SLPCR).  Orange bases depict the 5’ spliced leader and its primers. Blue bases depict the 3’ poly-A tail and its respective primers.   19	2.3 Results   2.3.1 Genes for plastid-targeted proteins obtained from a single-cell transcriptome.  Polykrikos lebouriae was identified by morphology in marine sand, and six single cells were manually isolated for transcriptome and genome sequencing (see below). The identification was confirmed by comparing DNA fragments of HSP90 and LSU rRNA genes from single cell sequence data to sequences obtained from previous isolates of Polykrikos lebouriae (LSU rDNA sequences shared 96.8% identity and HSP90 sequences shared 98.9% identity). In order to sequence the transcriptome, transcripts were reverse transcribed and amplified using dinoflagellate spliced leader and polyA primers.  For transcripts over 500 base pairs, the average length, after assembly, was 725 base pairs. Estimates of genome coverage were not possible as no sequenced genome is available for Polykrikos lebouriae or any species within its more inclusive clade (i.e., the Gymnodiniales).  SLPCR amplified an array of nuclear transcripts from P. lebouriae (Fig. 2.3), suggesting that the cell expressed genes spanning a broad range of functions, including photosynthesis. Fourteen transcripts over 600 base pairs long were associated with photosynthesis, and all were most closely related to dinoflagellates (Table 2.1). These were all nucleus-encoded, plastid-targeted genes, supporting the presence of a plastid that is functionally integrated.  Among these transcripts were two peridinin-chlorophyll a-binding precursor proteins, which are restricted to the peridinin-type plastids of dinoflagellates.         20	 2.3.2 Plastid-encoded genes obtained from single cell genomic data.  In order to examine the genome of the plastid itself, we also sequenced a genomic library created by multiple displacement amplification (MDA) from a single cell. Unlike the dinoflagellate specificity achieved through SLPCR, our total genomic amplification through MDA yielded a majority of reads (64%) from bacteria, with most of the remainder (34%) stemming from dinoflagellates, and a small fraction of viral or uncertain provenance (2%).  Of the eukaryotic reads, 5% were from plastids, with most other reads originating from the nuclear genome (which is generally quite massive in dinoflagellates).  Bacterial sequences were primarily from delta proteobacteria, specifically Francisella sp., which is known from cosmopolitan marine and freshwater strains as well as symbiotic strains found among animals and protists, existing either as parasites or commensal organisms (Schrallhammer et al. 2001). The eukaryotic sequences were most similar to dinoflagellates, as expected, and we identified and assembled three protein-coding genes from the plastid photosystem that are universally plastid-encoded: complete PsaA and PsbC genes and a partial plastid AtpA gene. After confirming the identity of each plastid photosystem gene using molecular phylogenetic analyses of the individual proteins  (Gavelis et al. 2015b, supplementary figures), the three proteins were concatenated and added to a 44-taxon alignment containing diverse dinoflagellates and other photosynthetic eukaryotes.  Both Bayesian analysis and maximum likelihood methods demonstrated that the P. lebouriae plastid sequences branch with homologues from peridinin-type plastids of other unarmored dinoflagellates (Fig. 2.4). The sequences from P. lebouriae were highly divergent, but branched with strong support after the Amphidinium clade and before the clade consisting of Togula jolla and all armored dinoflagellates.  Thus, the phylogenetic relationships inferred from the alignment of concatenated plastid-protein sequences are generally consistent with the placement of P. lebouriae as inferred from ribosomal gene sequences (Fig. 2.1).   21	  Figure 2.3  Transcripts expressed by a single cell isolate of Polykrikos lebouriae.  Transcripts are ranked from values 0 to 1 in abundance, and annotated according to Level 1 Subsystem hierarchical classification in MG-RAST. Predicted photosynthetic transcripts are shown in green.   22	 Figure 2.4.  Phylogenetic affinity of plastid genes in Polykrikos lebouriae. Maximum likelihood (ML) tree inferred from the 44-taxon alignment (1,595 unambiguously aligned amino acids) of concatenated plastid genes PsaA, PsbC, and AtpA using the PROTGAMMA model in RaxML.  Bootstrap support values 65 or higher and Bayesian posterior probabilities are listed above each branch.  The inset depicts a differential interference contrast (DIC) micrograph of the pseudocolony of Polykrikos lebouriae used for single-cell transcriptomics; this cell was undergoing mitosis when the image was captured (scale bar = 10 µm).    23	 Figure 2.5.  Peridinin-type plastid-targeting sequences. a. Class I transit peptides, each containing a transmembrane domain, which have been manually aligned, as have their “FVAP” motifs.  The average hydrophobicity score of each column in the transmembrane domain and neighboring regions have been plotted above the alignment. A: Fucoxanthin chlorophyll a/c-like protein. B: Ferredoxin. C: Light harvesting protein a. D: Light harvesting protein b. b. Class II transit peptides: These presequences lack a transmembrane domain, but contain the typical “FVAP” motif. E: ATP synthase gamma subunit. F: Acyl carrier protein. G: Photosystem II 12 kDa extrinsic protein. Amino acid color code: yellow=hydrophobic, blue=polar, green=negatively charged, red=positively charged.                  24	     Predicted Proteins # Top Hit E value Coverage Identity chloroplast ferredoxin 8 Alexandrium fundyense 6.00E-41 65% 68% chloroplast light harvesting complex protein 8 Symbiodinium sp.  1.00E-60 45% 61% chloroplast acyl carrier protein 5 Heterocapsa triquetra 2.00E-31 54% 65% plastid C1 class II fructose bisphosphate aldolase 4 H. triquetra 0 68% 85% chloroplast carbonic anhydrase 4 H. triquetra 7.00E-72 53% 71% chloroplast ATP synthase subunit C 4 A. affine 2.00E-32 60% 100% chloroplast phosphoribulokinase 3 Lingulodinium polyedrum 7.00E-164 93% 79% chloroplast ribose-5-phosphate isomerase 3 H. triquetra 5.00E-96 71% 69% chloroplast peridinin-chlorophyll a-binding protein precursor 2 A. tamarense 5.00 E-103 99% 82% chloroplast ATP synthase gamma subunit 2 H. triquetra 3.00E-125 87% 57% chloroplast ferredoxin-NADP{+) reductase 1 H. triquetra 1.00E-167 73% 70% chloroplast photosystem I, subunit III 1 H. triquetra 2.00E-42 43% 51% chloroplast photosystem II 12 kDa extrinsic protein 1 H. triquetra 7.00E-42 69% 62% chloroplast photosystem I subunit XI 1 H. triquetra 1.00E-85 74% 50%    Table 2.1. Nuclear-encoded plastid-targeted genes transcripts. Predicted plastid-targeted reads over 600 bp expressed by Polykrikos lebourae.   Identities were assigned using BLASTX queries against all proteins in Genbank.      25	2.4 Discussion   2.4.1 Utility of transcriptomics and genomics on single cells.  The majority of microbial species are not available in culture, and therefore the application of single cell methods at the genomic level is highly desirable. In this case, we used both single cell transcriptomics and single cell genomics to investigate the biology of plastids in P. lebouriae and test hypotheses for their origin, which were otherwise difficult to resolve. Single-cell spliced-leader transcriptomics was particularly powerful, and using this method we were able to obtain a diversity of nuclear-encoded transcripts from P. lebouriae, despite the presence of environmental contamination from bacteria (as seen in the MDA results) and potentially even other non-dinoflagellate eukaryotes.  Both nucleus-encoded transcripts and plastid-encoded genes consistently demonstrated that P. lebouriae is photosynthetic, with all photosynthesis related genes and transcripts grouping with those found in dinoflagellates with peridinin-type plastids, including a protein with the principle function of binding the pigment peridinin.  No abnormalities were seen in the plastid targeting sequences to suggest that P. lebouriae, which we found to bear typical Type I and II presequences (Fig. 2.5), as described in dinoflagellates with triple membrane bound peridinin plastids (Nassoury, Cappadocia, and Morse, 2003; Patron, Waller, Archibald and Keeling 2005). Thus is it unclear whether the two plastid membranes reported by Hoppenrath and Leander (2007a) were an accurate interpretation, a misinterpretation, or an artefact.   2.4.2 Peridinin plastids in Polykrikos lebouriae.  While we cannot falsify the possibility of transient or hidden plastids in some polykrikoids, our findings are contrary to the hypothesis that Polykrikos lebouriae acquired photosynthesis from diatoms or haptophytes and support the presence of peridinin-type plastids in P. lebouriae. The most parsimonious source for these  26	plastids is direct inheritance from ancestral polykrikoids.  Polykrikoid phylogeny, though lacking strong support at some deeper nodes, shows an unequivocal sisterhood between P. lebouriae and heterotrophic P. herdmaniae—which necessitates a recent loss of photosynthesis in P. herdmaniae.  A second loss is evident in the P. kofoidii - P. schwartzii clade, as they are strongly supported sister lineages, and therefore represent a loss independent from that found in P. herdmaniae.    2.4.3 What drove multiple losses of photosynthesis in polykrikoids?  Several losses of photosynthesis have previously been established in dinoflagellates (Saldarriaga, Taylor, Keeling, and Cavalier-Smith 2001), primarily among parasitic stem groups or within groups of questionable monophyly (e.g., the Gymnodiniales). Interestingly, multiple losses of photosynthesis appear to have occurred within polykrikoids alone, and the evolutionary reasons for this are worth considering. A prominent trend in polykrikoid evolution is a gradual increase in size (Hoppenrath and Leander 2007b) (Figure 2.1).  This makes polykrikoids effective predators, as they are able to consume dinoflagellates that are too large to be consumed by other predatory cells (Matsuoka, Cho, and Jacobson 2000).  Yet size is known to make photosynthesis less effective for single cells, as chloroplast self-shading increases, and absorptive surface area diminishes relative to volume. As a mixotroph, P. lebouriae is known to prey on other dinoflagellates (Aika Yamaguchi, personal communication), and our isolate possessed extrusive organelles such as nematocysts and taeniocysts.  The presence of such specialized predatory features, as well as mixotrophy and large cell size, may have predisposed polykrikoids to multiple losses of photosynthesis, as seen in P. herdmaniae, a sister species that shares the same habitat as P. lebouriae (Hoppenrath and Leander 2007a). Factors allowing the loss of photosynthesis probably vary by lineage, as losses have also occurred among smaller free-living and parasitic dinoflagellates.   27	 2.4.4 Future applications of SLPCR.  Understanding trends in the evolution of microbial eukaryotes will require a synthesis of ecology, phylogenetics, and genomics—the last of which has been particularly limited in its applications to uncultivated groups. While SLPCR has previously been applied to bulk RNA samples (Zhang et al. 2007, 2013) we show here that it is applicable to single cells.  In principle, this method is applicable to any system with uniform spliced leaders, as found in dinoflagellates, euglenids, kinetoplastids, and a growing number of invertebrates. SLPCR shows promise not only in avoiding contaminants in environmental isolates, but in capturing gene expression of a single cell at a given point in time, such as stages of the cell cycle, cells perturbed by experimental stimuli, or simply cells in the dynamism of their natural habitats.   28	3 Eye-like ocelloids in warnowiids are built from different endosymbiotically acquired components.   3.1 Introduction  Many organisms can orient to light. In some single-celled eukaryotes, such as Chlamydomonas and many dinoflagellates, an “eyespot” directs photons onto photoreceptors on the flagellum, allowing the cell to respond to the intensity and direction of light (Foster and Smyth 1980). A vastly more complex structure is found in warnowiid dinoflagellates: the eye-like ocelloid (Fig. 3.1).  Ocelloids consist of subcellular components resembling a lens, a cornea, iris-like rings and a pigmented cup called the retinal body (Greuet 1970), which together so resemble the camera-type eyes of some animals that they have been speculated to be homologous (Gehring 2004).  The first description of a warnowiid was dismissed as a cell that had scavenged the eye from a jellyfish (recounted in Kofoid and Swezy 1921). Ultrastructural studies of the ocelloid subsequently suggested that the retinal body might be derived from a plastid, in that it contains thylakoid-like membranes during cell division (Greuet 1976).   The ocelloid is among the most complex subcellular structures known, but its function and evolutionary relationship to other organelles remains unclear (Hoppenrath et al. 2009.) This poor state of knowledge can be attributed to the fact that warnowiids are uncultivated and rarely encountered in environmental samples, with as few as two cells reported from the plankton per year for some species (Gomez 2009). Modern single-cell genomics and microscopy approaches, however, provide opportunities to study uncultivated eukaryotes at the molecular and ultrastructural levels, including rare species (Kolisko et al. 2014, Gavelis et al. 2015b). In an attempt to learn more about the cell biology of ocelloids, we applied single-cell transcriptomics on two genera of warnowiids: Erythropsidinium sp., and Warnowia sp., as well as TEM on Erythropsidinium and Nematodinium sp. Lastly, we investigated the 3-dimensional ultrastructure  29	and phylogenetic origin of the retinal body in Nematodinium sp. by using focused ion beam scanning electron microscopy (FIB-SEM) on isolated cells, and single-organelle genomics.   Figure 3.1 Light micrographs (LM) of warnowiids used in this study. a. Still frame LM from a video of Warnowia sp. b. LM of Erythropsidinium sp. c. LM of Nematodinium sp. with a nematocyst (arrowhead). d. LM of the ventral side of Nematodinium sp. showing red pigmentation of the retinal body. e. Epifluorescence image of the same cell and angle, showing red fluorescence of the retinal body excited by 505 nm light. f. LM of Nematodinium sp. showing a bright spot of reflectivity (i.e., “eyeshine”) (arrowhead) in the ocelloid.   				 30	3.2 Materials and methods   3.2.1 Collection of organisms.  From 2005 to 2009, Erythropsidinium sp. and Warnowia sp. were collected from the marine water column in Suruga Bay, Japan (34.929722, 138.400262). On an inverted light microscope, cells of Warnowia sp. were recognized as ocelloid-bearing cells encircled three or more times by a helical groove (Fig. 3.1a). Cells of Erythropsidinium sp. were identified based on the presence of an ocelloid and a piston organelle (Fig. 3.1b). cDNA libraries from four pooled cells of Warnowia sp. and two pooled cells of Erythropsidinium sp. were prepared as described in section 3.2.4. In the summer of 2012 and 2013, Nematodinium sp. was collected from surface water in Bamfield Inlet, Bamfield, BC, Canada (48.834354, -125.136929) with a 20 µm plankton net. Cells of Nematodinium sp. were identified based on the presence of an ocelloid and nematocysts (Fig. 3.1c, d). Uncultivated Nematodinium sp. cells containing putative prey organisms (visible as pigmented vacuoles) were chosen for transmission electron microscopy, so that their feeding habits could be inferred from intracellular remnants.  In total, 12 cells of Nematodinium sp. were fixed and mounted individually for TEM, and 58 cells of Erythropsidinium sp. were obtained and mounted for TEM in groups.    3.2.2 Fluorescence and DIC microscopy.   Red epifluorescence of the Nematodinium sp. retinal body was excited with a 505 nm Argon laser on a Zeiss Axioplan inverted light microscope (Fig. 3.1 e).  Differential interference contrast (DIC) observations of Nematodinium sp., Warnowia sp., and Erythropsidinium sp., were performed using the same microscope.    31	 3.2.3 Single-cell transmission electron microscopy of uncultivated Nematodinium sp.  Each isolated cell of Nematodinium sp. was micropipetted onto a poly-L-lysine coated slide. Cells were fixed with 2% glutaraldehyde in filtered seawater for 30m on ice.  After two washes in filtered seawater, cells were post-fixed in 1% OsO4 for 30m. Cells were dehydrated through a graded series of ethanol (50%, 70%, 85%, 90%, 95%, 100%, 100%) at 10m each, and infiltrated with a 1:1 acetone-resin mixture for 10m.  Cells were infiltrated in Epon 812 resin for 12h, after which the resin was polymerized at 60° C for 24h to produce a resin-embedded cell affixed to the glass slide. Using a power drill, resin was shaved to a 1 mm3 block, which was removed from the glass slide with a fine razor. The block, containing a single cell, was attached with Super Glue to a resin stub in the desired orientation for sectioning.  Thin (45 nm) sections were produced with a diamond knife, post-stained with uranyl acetate and lead citrate and viewed under a Hitachi H7600 Transmission Electron Microscope.  3.2.4 Isolation of the retinal bodies of Nematodinium sp.  In preparation for single-organelle genomics, five cells of Nematodinium sp. with no visible prey contents were selected in order to minimize the chances of genetic contamination.  Each cell of Nematodinium was micropipetted onto a slide in a droplet of TE buffer and affixed to a patch of poly-L-lysine.  Cells were lysed with nuclease-free water.  The nucleus and other cell contents were gently dislodged with rinses of TE buffer, leaving the retinal body behind for manual isolation. Unlike the retinal body, which is darkly pigmented, the cornea and mitochondria of the ocelloid are much smaller, transparent and could not be isolated after cell lysis or tracked through rinse steps.  Five different retinal bodies were isolated and pooled onto a new, sterile slide, and washed three times with TE buffer to remove as many other cellular remnants as possible.      32	 3.2.5 Single-organelle genomics of Nematodinium sp.  In order to test for the presence of a plastid genome in the retinal body, we performed a genomic amplification using phiX 29 polymerase (Repli-G mini kit, Qiagen) on five individually isolated retinal bodies that were then pooled together.  We performed a control reaction by amplifying a pool of five whole cells of Nematodinium sp. using the same procedures as for the retinal bodies. The whole cell amplification provided a measure of overall plastid DNA concentration, against which the retinal body plastid DNA concentration could be compared.  In order to minimize amplification bias, each reaction was divided into four aliquots, run in parallel, and pooled after the 15h amplification period. Paired end sequencing on an Illumina MiSeq yielded 9,798 reads from the retinal bodies, versus 501,338 reads from whole cells. From these reads, plastid genes were assembled using the de novo assembly program Ray (Boisvert, Laviolette, and Corbeil 2010) which fragmented the reads into a variety of hash sizes (“kmers”), then assembled them.  We found the assembly from 53 bp kmers to be optimal, as they produced the longest contigs, including six partial plastid genes. In order to estimate the concentration of plastid reads in the whole cell vs. isolated retinal body amplifications, we counted plastid reads in Bowtie (Langmead, Trapnell and Salzberg 2009), a read mapping program, then divided them by the total number of reads sequenced from that reaction.    3.2.6 Molecular phylogenetic analyses.  The six plastid genes, photosystem I P700 apoprotein A2 (PsaB), photosystem II protein D1 (PsbA), photosystem II CP47 protein (PsbB), photosystem II protein D1 (PsbD), cytochrome b6 (Petb), and cytochrome b6/f complex subunit 4 (PetD) were translated, and their amino acids aligned with a representative set of eukaryotes in Muscle (Edgar 2004) with fast-evolving and ambiguously aligned regions removed in Gblocks 0.91b (Castresana 2000). The amino acid substitution model (Protein GTR gamma) was estimated from the concatenated  33	alignment of 1,618 amino acids using the Models package in Mega 6.0.5 (Tamura et al. 2013). A maximum likelihood phylogeny was run with 500 bootstraps in RAxML (Stamatakis 2006). A second, Bayesian analysis was run for 10,000 generations in MrBayes 3.2 (Ronquist and Huelsenbeck 2003), using the high-heating setting of (nchains = 4), to account for rapid evolution of dinoflagellate plastids.  These maximum likelihood analyses were run both for the multiprotein dataset, and also for each protein individually. A dinoflagellate phylogeny was estimated using SSU and LSU ribosomal rDNA sequences, concatenated as a 2,331 nucleotide alignment, across 36 dinoflagellate taxa including published sequences from Nematodinium sp., Warnowia sp. and Erythropsidinium sp.  3.2.6 Focused ion beam scanning electron microscopy (FIB-SEM).  Cells of Nematodinium sp. were individually transferred into a droplet of 20% bovine serum albumin in phosphate buffered saline solution (an isotonic solution in which cells are less likely to lyse or become distorted).  Cells were frozen immediately to minimize fixation artefacts, using a Leica EM HPM 100 high-pressure freezer (Leica, Wetzlar, Germany).  Freeze substitution was subsequently used to remove the aqueous content of the cells and replace it with an acetone solution containing 5% water, 1% osmium tetroxide and 0.1% uranyl acetate, at -80 °C for 48h, -20 °C for 6h, then transitioned back to 4 °C over 13 h.  The prepared samples were washed twice in 100% acetone.  Two cells were recovered by micropipette.  Each cell was placed on a separate Thermonox (Fahlenbach, Germany) coverslip, where it adhered to a patch of poly-L-lysine.  In preparation for FIB-SEM, cells were infiltrated with a 1:1 mix of acetone and Embed 812 resin for 2h, then 100% resin overnight.  A second Thermonox coverslip was applied, sandwiching each cell in a thin layer of resin between the coverslips. Resin was polymerized at 65 °C for 24h. The top coverslip was then removed with a razor blade to expose the resin face overlying the cell.     34	 A single cell was imaged by a FEI Helios NanoLab 650 dual beam FIB-SEM.  The ion beam milled through the cell in 20 nm increments, yielding 190 image slices. Slices were aligned as a z-stack in Amira 5.5.  Features of interest, including mitochondria and chloroplasts, were semi-automatically segmented: that is, manually traced in approximately one of every three slices, before automatic interpolation filled in the volumes between the slices. Images that did not pass quality screening because of fluctuations in microscope beam power and autofocus were not directly segmented, but were interpolated from segmentation on neighboring images, per manufacturer’s instructions. Surfaces of the mitochondria, chloroplasts, and vesicles were generated, smoothed and colorized to produce a 3-dimensional model of the components that form the ocelloid.   3.3 Results  3.3.1. TEM in Nematodinium sp. and Erythropsidinium sp.  Thylakoid-like structures have been reported only once before in the retinal body (Greuet 1979), so we examined the ultrastructure of the ocelloid in Nematodinium sp. and Erythropsidinium sp. using single-cell transmission electron microscopy (TEM). During interphase, the retinal body contains highly ordered waveform membranes (Fig. 3.2), which are perpendicular to the plane expected for thylakoids in a chloroplast. However, we confirmed that near the end of interphase, the waveform membranes dedifferentiated into a plastid-like arrangement made of double-stacked thylakoid-like structures (Figs. 3.3c, 3.4d, and 3.5). Thus, the thylakoids and waveform membranes represent two modes of the same membrane system. Moreover, we found that the retinal body of Nematodinium sp. exhibits red fluorescence under 528 nm (green) light—suggesting the presence of chlorophyll or another autofluorescent pigment (Fig.  35	3.1e).  In Nematodinium, we also found mitochondria in the ocelloid, where they formed a cornea-like layer overlying the lens (Fig. 3.6c) (Greuet 1977).    36	Figure 3.2 Ultrastructure of the retinal body in Nematodinium sp. A composite of 12 electron micrographs showing a glancing section through the retinal body, which is contains stacked waveform membranes (white square and inset) enveloped by pigmented lipid droplets (*).    Figure 3.3 Development in warnowiids a., b. Light micrographs of a single cell of Nematodinium sp., and single cell of Erythropsidinium sp., progressing from interphase (left) to division (right). Scale bars = 10 µm.  c.  Transmission electron micrographs of membranes in the retinal body from cells at different stages of development, including differentiated (left, Nematodinium sp.), transitional (middle, Erythropsidinium sp.), and dedifferentiated modes (right, Nematodinium sp.).  Scale bars = 200 nm. The double arrowhead marks a typical plastid; arrowheads mark the retinal bodies; arrows mark lenses that are dedifferentiating.  37	 Figure 3.4 Thylakoid forms in Nematodinium sp. a. TEM showing a small, peripheral plastid in Nematodinium sp. with typical thylakoids resembling peridinin plastids in other dinoflagellates. b. TEM showing thylakoids in the iris region of the ocelloid. c. TEM showing thylakoids in the pigmented iris (p) beside waveform membranes of the retinal body (r), during interphase. d. TEM showing a retinal body towards the end of interphase, in which the waveform membranes (w) dedifferentiate and are continuous with the typical thylakoids (arrows).    38	 Figure 3.5 Transient thylakoids in the retinal body as viewed with transmission electron microscopy. a, b. Ocelloid in a cell of Nematodinium sp. near division. c, d.  Ocelloid in a cell of Erythropsidinium sp. during division. L = lens; t = thylakoids; asterisks = lipid droplets; arrows = waveform membranes.    39	3.3.2 Transcriptomics in Erythropsidinium sp. and Warnowia sp.  From polyadenylated cDNA libraries of Erythropsidinium sp. and Warnowia sp., we found that these heterotrophic genera expressed multiple photosynthesis-related genes (Table 3.1), including light-harvesting proteins. In addition, Warnowia sp. expressed three transcripts corresponding to the chloroplast-soluble peridinin-chlorophyll-binding protein, which is distinctive for dinoflagellate peridinin-type plastids.  3.3.3 Single-organelle genomics in Nematodinium sp.   In order to investigate the phylogenetic origin of the retinal body more directly, we characterized genes encoded on DNA associated with the organelle structures. Single cells of Nematodinium sp. were micro-dissected, and individual retinal bodies were isolated (Fig 3.6. In order to compare the DNA content of dissected organelles to the DNA content of whole Nematodinium cells (including nuclei), we also pooled five intact Nematodinium sp. cells and subjected them to the same procedures for DNA amplification and sequencing. From sequence databases derived from both samples, we identified genes that are encoded in the plastid of other dinoflagellates. Overall, six plastid genes were identified from isolated retinal bodies, PsaB, PsbA, PsbB, PsbD, PetB, and PetD, spanning photosystems I and II.  These genes grouped strongly with the peridinin-containing plastids of dinoflagellates in individual and concatenated phylogenetic analysis (Fig. 3.7), and collectively, plastid-encoded genes represented 13% of all reads (Fig. 3.6d). By contrast, the proportion of plastid/nuclear DNA in the whole-cell amplification was <0.0001% (Fig. 3.6e). The representation of plastid DNA in the retinal body was, therefore, over 1,600 fold higher than in whole cells.  40	 Figure 3.6 Genomics and structure of organelles in the ocelloid. a. Illustration of Nematodinium showing the basic components of the ocelloid with their putative organellar origins.  b.  Transmission electron micrograph (TEM) of the ocelloid of Erythropsidinium, including the lens (L) and retinal body (r). c. TEM of the ocelloid of Nematodinium, depicting the edge of the lens (L) where it is overlain by a cornea-like layer of mitochondria (m). d. Genomic reads amplified from five whole cells of Nematodinium; arrow, retinal body. e. Genomic reads amplified from five retinal bodies (arrow) after they were micro-dissected from individual cells of Nematodinium.    41	 Figure 3.7 Phylogeny of retinal-body encoded proteins. Six partial plastid genes from the retinal body of the ocelloid in Nematodinium sp. were amplified. Photosystem I P700 apoprotein A2, photosystem II protein D1, photosystem II CP47 protein, photosystem II protein D1, cytochrome b6, and cytochrome b6/f complex subunit 4 were translated and concatenated for a 1,618 amino acid alignment. The tree was inferred by analyzing the 42-taxon alignment using maximum likelihood (ML). Statistical support for the branches was evaluated using 500 ML bootstrap replicates and Bayesian posterior probabilities.  Support values are shown for all branches within the Myzozoa (dinoflagellates and chromerids).    42	  3.3.4 Three-dimensional reconstruction of the ocelloid in Nematodinium sp.   To investigate the physical connections between the different components of the ocelloid and surrounding structures, such as peridinin-type plastids, we performed FIB-SEM tomography on a single isolated cell of Nematodinium sp. The three-dimensional reconstructions of our FIB-SEM data demonstrated that the outer membrane of the retinal body is fused to a network of adjacent plastids, forming a membranous web throughout the cell (Fig. 3.8). Therefore, the retinal body appears to be a differentiated region of a larger, netlike plastid.    Tomographic reconstructions also confirmed a close association between mitochondria and the lens of the ocelloid.  The mitochondria surrounding the lens were interconnected and formed a sheet-like “cornea” layer consistent with TEM data. The corneal layer surrounded all regions of the lens except for a few minor perforations and the side facing the retinal body (Fig. 3.8, 3.9). The corneal mitochondria appear to form a continuous network with mitochondria in the nearby cytoplasm. The ocelloid, therefore, represents an intriguing mixture of components with endogenous and endosymbiotic origins.            43	  Figure 3.8 Three-dimensional reconstruction of the ocelloid of Nematodinium sp. using FIB-SEM tomography. a. Stack of a halved cell, showing the nucleus and the ocelloid (box). b. FIB-SEM slice of the ocelloid. c. Translucent FIB-SEM stack of the region surrounding the ocelloid, including the lens (yellow) and full plastid network (red). d. Reconstructions of the ocelloid and its component parts, including the mitochondrial cornea-like layer, vesicular lens, and retinal body. Color coded organelles: blue = mitochondria; yellow = lens; red = plastids.    44	 Figure 3.9 FIB-SEM slices of the cornea-like layer of mitochondria in the ocelloid of Nematodinium sp. a. Low-magnification TEM of the ocelloid, with rectangles delimiting the areas of higher magnification shown in images b, c and d.  b-d. High magnifications of structures bordering the lens (L). Mitochondria, m; pigmented ring, p; retinal body, r.    45	 Figure 3.10 Inferences on the predatory nature of Nematodinium sp. from live and preserved specimens. a. Differential interference contrast LM showing a cell with prey (P) visible as green tinted food vacuole. b. Differential interference contrast LM showing a cell in which the condensed dinoflagellate-type nuclei (n) are visible as birefringent chromosomes in both the predator and prey. c. Differential interference contrast LM of a Nematodinium sp. cell containing digested prey (arrowhead) and co-occurring with potential prey, a smaller dinoflagellate. d. TEM showing a food vacuole inclusion consisting of a bolus of discharged trichocysts. e. TEM of undischarged dinoflagellate-type trichocysts showing their characteristic square shape in transverse section. f. TEM of discharged dinoflagellate-type trichocysts showing their characteristic striation pattern in longitudinal section.  46	3.4 Discussion  Prior to this study, there was little evidence for homology between the ocelloid and other structures found in dinoflagellates.  Based on its resemblance to camera-type eyes, a relationship was even suggested between the ocelloid and the eyes of some animals (Gehring 2004). To the contrary, our findings indicate that the ocelloid is a conglomerate of several membrane-bound organelles, including endomembrane vesicles, mitochondria and plastids.  The ocelloid is likely homologous to the much simpler eyespots found in several other lineages of dinoflagellates, most of which share features in common with the peridinin plastid (Dodge 1984). Peridinin plastids stem from an ancient red alga that was incorporated by the common ancestor of all myzozoans (dinoflagellates, chromerids, and apicomplexans), many of which (including all apicomplexans) subsequently lost photosynthesis and reduced their plastids to cryptic, morphologically simple structures (Janouskovec et al. 2010). While morphological reduction is a common trend among endosymbiotic organelles, the ocelloid in warnowiids demonstrates that increased complexity can also arise.   Still, virtually nothing is known of ocelloid function, due to a lack of recorded observations on live warnowiids. This is because warnowiid cells are rarely encountered, have never been cultivated, and degrade rapidly when removed from the plankton (Gomez 2009). Nevertheless, we observed one important detail of warnowiid life history using TEM of individual cells isolated directly from the ocean. We found that the food vacuoles in Nematodinium contained trichocysts (Fig. 3.10d-f), which are defensive extrusive organelles found in dinoflagellates. Given the presence of dinoflagellate remnants in its food vacuoles, it is logical to conclude that Nematodinium preys on other dinoflagellates.   Our finding that Nematodinium preys on other dinoflagellates opens the way for some interesting—though as yet unsupported—speculation about the ocelloid’s  47	function. Perhaps the ocelloid is involved in the detection of other dinoflagellates as prey. Some dinoflagellates are capable of bioluminescence (Widder 2010), which may be what ocelloids detect, but all dinoflagellates contain a distinctively large nucleus of permanently condensed chromosomes, and these chromosomes polarize light (Liu and Kattawar 2013). An intriguing possibility is that the ocelloid can detect polarized light, and by extension, dinoflagellate prey. The structure of the retinal body seems to provide some support for its function as a polarization filter, since its waveform membranes could potentially screen out non-polarized light.   Testing such a specific phototactic behavior will be challenging until warnowiids are brought into culture. This would allow for live observation or warnowiid behavior (including hunting), as well as testing for whether cell exhibit polarotaxis in the presence of circularly polarized light. Nevertheless, the genomic and detailed ultrastructural data presented here have resolved the basic components of the ocelloid and their origins, and demonstrate how evolutionary plasticity of mitochondria and plastids can generate an extreme level of subcellular complexity.    48	4 Comparative ultrastructure and genomics suggest that nematocysts evolved at least twice.  4.1 Introduction  Projectile organelles (extrusomes) are used by many groups to either capture or repel other organisms, though it is unclear how, or how many times, extrusomes evolved. We examined the harpoonlike extrusomes called “nematocysts” in two different groups of dinoflagellates (Figure 4.1), which are among the most complex organelles described. The nematocysts of Polykrikos are often compared to those of cnidarians, which likewise discharge a coiled tubule from within a capsule (Holstein 1981)—and these structures have been suggested to be homologous (Hwang et al. 2008, Shostak 1993, David et al. 2008). Complex nematocysts have also been described in some ciliates (Raikov 1992), and an increasing number of dinoflagellates (e.g., Nematodinium sp., Paragymnodinium, Gyrodiniellum (Greuet 1987; Kang et al. 2010, 2011). In ultrastructural studies (Westfall, Bradbury, and Townsend 1983; Mornin and Francis 1979; Greuet 1977, 1989), the nematocysts in dinoflagellates have been interpreted as containing stylets and opercula—features also found in cnidarians (Holstein 1981).  Because cnidarians and dinoflagellates are separated by a vast expanse of lineages without extrusomes (Figure 4.2), they probably evolved these ballistic organelles independently, in which case any supposed similarities would have resulted from convergent evolution (Leander 2008a). However, a recent study indicated that dinoflagellate nematocysts also contain minicollagens (Hwang et al. 2008), which are the principle structural proteins of nematocysts in cnidarians (Thomas et al. 2008). This recent finding appeared to legitimize a hypothesis in which nematocysts were disseminated via lateral gene transfer (Shostak 1993).  Specifically, Shostak hypothesized that such a transfer occurred during an  49	ancient dinoflagellate-cnidarian symbiosis (i.e., between a basal anthozoan and an ancestral Symbiodinium like-dinoflagellate)—and this symbiogenic hypothesis has since been reiterated in different forms (Hwang et al. 2008, Thomas et al. 2008). Here we test these hypotheses by (1) comparing genomic, proteomic and transcriptomic data from dinoflagellates and cnidarians and (2) establishing detailed 3-D ultrastructural models of dinoflagellate nematocysts using high resolution microscopy (e.g., focused ion beam scanning electron microscopy). 	 50	 Figure 4.1 Nematocysts in Polykrikos kofoidii and Nematodinium sp. a. Polykrikoid kofoidii. b. Nematodinium sp., with the ocelloid positioned out of view on the opposite side of the cell. c. A discharged nematocyst from a crushed cell of P. kofoidii. Scale bars = 10 µm. 	 51	 Figure 4.2 Distribution of extrusomes in eukaryotes.  a. – e. Illustrations of extrusive organelles with a coiled thread/tube. a. Nematocyst of Polykrikos spp. b.  Toxicyst of colponemids. c. Nematocyst of the ciliate Remanella rugosa (Raikov 1992). d. “Long toxicyst” of the ciliate Loxodes sp. e. Nematocyst or “cnidocyst” of cnidarians (an atrichous isorhiza). Gray lines in the phylogeny represent poorly supported branches. Taxa colored with red have extrusomes that eject a tube/thread from within a pressurized capsule (e.g., toxicysts or nematocysts). Taxa colored with blue have extrusomes consisting of an expanding lattice (e.g., trichocysts and mucocysts). Taxa colored with green have extrusomes that consist of uncoiled tubes (e.g., ejectisomes). For clarity, whole-cell ballisitics (e.g., microsporidians, helicosporidians, and Haptoglossa) are not included. Phylogenetic tree based on Burki 2014.       52	4.2 Materials and methods  4.2.1 Genome and transcriptome mining for nematogenic proteins.   A proteomic dataset from nematocysts in Hydra magnipapillata was provided by Thomas Holstein and Suat Ozbek as a reference to search for nematogenic protein genes within the genomes of other eukaryotes. I used BLASTP to query against a custom database that I assembled from published proteomes (predicted from high-coverage genomes) from a representative group of 27 eukaryotes (Figure 4.3). I also used tBLASTn to search against the transcriptomes that we previously obtained from Polykrikos lebouriae (in Chapter 2), as well as a public transcriptome from the myxozoan Tetracapsuloides bryosalmonae to compensate for the low coverage of the only available myxozoan genome (from Thelohanellus kitaui). Together these 29 taxa included: four cnidarians (two with nematocysts and two with polar filaments); an extrusome-bearing cryptophyte (Guillardia theta, which has ejectisomes); an extrusome-bearing chlorarachniophyte (Bigelowiella natans); two ciliates with extrusive mucocysts or trichocysts (Tetrahymena thermophila and Paramecium tetraurelia); two dinoflagellates with extrusomes (Symbiodinium microadriaticum with mucocysts, and Polykrikos lebouriae with mucocysts, trichocysts, taeniocysts, and nematocysts); two parasites that lack extrusive organelles but undergo ballistic sporulation (Encephalitozoon cunuculi and Helicosporidium sp.); and two apicomplexans that have complex, non-ballistic secretory organelles (Cryptosporidium hominis and Toxoplasma gondii). To our knowledge, these represent all available genomes from protists with extrusomes or other ballistic structures.  Genomes were BLAST queried using default parameters (E value threshold: e-10). For each query, the top hit was reciprocally queried using the BLASTP algorithm of the NCBI BLAST server (http://blast.ncbi.nlm.nih.gov/Blast.cgi) against the ‘nr’ database to investigate whether these proteins shared conserved  53	domains and if they constituted best reciprocal BLAST hits—a common metric for predicting gene orthology  (Figure 4.2).   4.2.2 Collection of organisms.  In the spring and fall of 2011 to 2015, cells were collected with a plankton net (20 µm mesh) at two sites in British Columbia, Canada: a dock at Granville Island, Vancouver (49.271337, -123.137747), and a pier off the sea-plane port on Patricia Bay, North Saanich (48.653266, -123.448652). Using a Zeiss Axioplan inverted light microscope, cells of Nematodinium sp. were identified based on the presence of an ocelloid and nematocysts (Fig. 3.1c, d), and cells of Polykrikos kofoidii were identified based on the absence of plastids and the presence of two nuclei and eight flagellated segments (a.k.a. “zooids”) per pseudocolony. Over this time period, approximately 12 cells of Nematodinium sp. were fixed and mounted individually for TEM; approximately 20 cells of Polykrikos kofoidii were obtained and mounted for TEM either individually or in groups; and 10 more cells of P. kofoidii were crushed and lysed to allow for SEM of isolated extrusomes.   4.2.3 Inducing nematocyst discharge.   Using the methods of Hovasse (1951), pseudocolonies of Polykrikos kofoidii were crushed on slides to induce nematocyst discharge (i.e., by striking the center of the coverslip with the butt of a plastic screwdriver, quickly to avoid grinding).  Approximately a third of the nematocysts in each preparation were discharged. To remove the coverslip without smearing the nematocysts on the slide beneath it, distilled water was slowly added to the seam where the slide and coverslip met, allowing the coverslip to gently float off the surface (nematocysts generally adhered to the slide).  Once the coverslip was removed, the slide and nematocysts were rinsed three times with distilled water to remove lysed cell debris. Currently no method is known to discharge the nematocysts of Nematodinium sp.—nor has any firing event been described; however, I  54	attempted to stimulate discharge using (1) the striking method of Hovasse (1951) and (2) Bouin’s fixative, which was used successfully by Hwang et al. (2008) on polykrikoids. These techniques were unsuccessful in Nematodinium sp., as was exposure of live slide-mounted cells to sonication, 9 volts of electricity, and potential prey dinoflagellates, Scrippsiella trochoidea and Heterocapsa triquetra.   However, I did see a nematocyst discharge on one occasion, from a cell of Nematodinium sp. that had been slightly agitated under a coverslip; unfortunately, the slide preparation was crushed when I attempted to capture an image under oil immersion. Nevertheless, I saw a capsule launch out of the cell at a distance of approximately 15 µm and adhere to the coverslip by its tapered tip (as in polykrikoids). The capsule remained attached to the cell by some element that I could not discern, but was evidently tethered because water currents could not drag the cell more than 15 µm from the capsule, and at this radius, the cell swung in rotation around where the capsule had adhered to the coverslip. This suggests that nematocysts in both Nematodinium sp. and polykrikoids behave in a similar harpoon-like manner—though this observation clearly needs to be verified in detail.   4.2.4 Scanning electron microscopy on discharged nematocysts.   Slides with extruded nematocysts from Polykrikos kofoidii were dehydrated through a graded series of ethanol washes (50%, 70%, 85%, 90%, 95%, 100%, 100%) at 10 min each, after which the ethanol was sublimated in an Autosamdri 815A critical point dryer. The specimens were sputter coated with 5 nm of gold coating before being imaged in a Hitachi S4700 scanning electron microscope.   4.2.5 Single-cell transmission electron microscopy.  Each isolated cell of Polykrikos kofoidii was micropipetted onto a poly-L-lysine coated slide. For 30 min on ice, cells were simultaneously fixed/post-fixed in a  55	solution of 2% glutaraldehyde and 1% OsO4 in filtered seawater. Fixative was removed with two rinses of distilled water, after which a droplet of warm low-melting point agar was dropped onto each cell, sealing it onto the slide and preventing the cell from being lost later in preparation. Cells were then dehydrated through a graded series of ethanol (50%, 70%, 85%, 90%, 95%, 100%, 100%) at 10 min each, then through two rinses of 100% acetone.    Cells of Nematodinium sp. were fixed using an improved high-pressure freezing and freeze-substitution method. Each cell was transferred by mouth pipet into a droplet of 20% bovine serum albumin in 1X phosphate buffered saline (an isotonic solution less likely to distend or lyse the cell). Cells were frozen immediately to minimize fixation artefacts, using a Leica EM HPM 100 high-pressure freezer (Leica, Wetzlar, Germany). Freeze substitution was subsequently used to remove the aqueous content of the cells and replace it with an acetone solution containing 5% water, 1% osmium tetroxide and 0.1% uranyl acetate, at -80 °C for 48 h, -20 °C for 6 h, then graded back to 4 °C over 13 h. The prepared samples were washed twice in 100% acetone. Two cells were recovered by micropipette. Each cell was placed on a separate Thermonox (Fahlenbach, Germany) coverslip, where it adhered to a patch of poly-L-lysine.   Once in 100% acetone, cells of both Polykrikos lebouriae and Nematodinium sp. were infiltrated with a 1:1 mix of acetone and Embed 812 resin for 2 h, then 100% resin overnight, after which fresh resin was added polymerized at 60 °C for 24 h. This produced a resin-embedded cell affixed to the Thermonox slide. Further prep for FIB-SEM was described in section 3.2.6 and 3.3.4, while TEM sections were sectioned on into 45 nm sections with a diamond knife, and post-stained with uranyl acetate and lead citrate for viewing under a Hitachi H7600 TEM.     56	4.3 Results   4.3.1 No predicted dinoflagellate proteins with affinity to cnidarian nematocyst machinery.   Searches of known cnidarian nematocyst proteins (e.g. minicollagens, spinalin, cnidoin) did not return best reciprocal-BLAST hits (an established method for predicting gene orthology), aside from those already found in cnidarians. Several opisthokonts exhibited proteins with moderate similarity to minicollagens and nematogalectin, as they shared collagen domains; but no collagens were found in clades diverging before sponges. Some weak protist hits were found to nematogalectin, but while they shared a lectin domain (which is common across archaea, bacteria and eukaryotes), these hits did not constitute paralogs, for they lacked collagen domains as well as cysteine-rich regions necessary for polymerization. Other known nematogenic proteins do not exhibit significant hits.   4.3.2 SEM and TEM illustrate a lack of homology between polykrikoid and cnidarian nematocysts  Our ultrastructural analysis of nematocysts in Polykrikos kofoidii challenges previous interpretations of cnidarian-like structures in dinoflagellate nematocysts. While central elements in the nematocysts of both polykrikoids and Nematodinium sp. have typically been interpreted as “stylets,” we found no evidence for any such structural or functional homology. When viewing nematocyst discharge in Polykrikos kofoidii for the first time through SEM, it was evident that P. kofoidii does not discharge a stylet as in cnidarians (Figure 4.4). TEM images of undischarged nematocysts show that the “stylet” is simply a portion of the tubule that passes through a pore in the anterior chamber wall (Figure 4.4l).  Thus, its previous labeling as a “stylet” was probably an over-interpretation inspired by the use of cnidarian terminology (i.e., by naming dinoflagellate extrusomes as “nematocysts”). Likewise, the complex cap in  57	polykrikoid nematocysts has generally been called an “operculum” (and I continue to use the term here for clarity only), but there is no evidence that it opens in the hatch-like manner as found in cnidarians (Figure 4.4). Rather, the nematocyst tubule exits through a small pore in the capsule (Figure 4.4e).  However, we did notice a new feature in the nematocysts of P. kofoidii, which it appears to share with those of Nematodinium sp. Though faint, a characteristic banding pattern was seen in the capsule wall of P. kofoidii, and these striations strongly suggested a homology to the material of the capsule wall in Nematodinium sp., which has striations that likewise repeat in 45 nm intervals (Figure 4.4k). This represents the most compelling evidence of homology between the highly divergent extrusomes of polykrikoids and Nematodinium sp. Ideally, this would be confirmed by proteomics as well—though these methods are not available for uncultivated cells like warnowiids.   4.3.3 TEM and FIB-SEM reveals new levels of complexity in the nematocysts of Nematodinium sp. 	High pressure freezing and freeze-substitution allowed for better fixation than has been attained previously for the nematocysts of Nematodinium sp, and, as such, we were able to identify several new structures. Rather than being capped by a globular, indistinct “operculum,” as was previously described (Mornin and Francis 1976; Westfall 1987), the top of the capsule is overlain by a “distal rosette.” The rosette has radial symmetry (unlike any known cnidarian nematocyst), and is divided into several identical “petals.” Each petal overlies a structure that we call a “plug.” We interpret plugs as the equivalents of opercula, because each plug covers an opening to the capsule below.  Remarkably, the nematocysts varied between 11 and 14 fold symmetry (with just as many plugs, petals, and capsules). Nematocysts of Nematodinium sp. have been previously reported to have 7 capsules (by Greuet) or 14 (by Mornin) but  58	neither author appeared to notice that these symmetry numbers differed between their studies. These biological differences might not be surprising, given that each author collected from a different ecosystem: in the Mediterranean and Pacific Ocean, respectively.  But our description of high local variation seems to suggest that we have underestimated the variability of these complex extrusomes.   59	 Figure 4.3 Distribution of cnidocyst machinery in eukaryotes. Predicted proteins from the nematocyst proteome of Hydra magnipapillata (a cnidarian) were blasted against proteins predicted from the genomes  (solid boxes) or transcriptomes (slashed boxes) of the listed eukaryotic species. Best reciprocal blast hits are designated by white boxes—and are predicted to be orthologs.  Black and gray boxes are predicted to have no orthology. Groups in red squares have “nematocysts,” based on morphological descriptions.  60	  Figure 4.4 Nematocysts in Polykrikos kofoidii. a.—f. SEMs of undischarged (a., b.), partially discharged (e., f., c.) and fully discharged (d.) nematocysts isolated from crushed cells. e. and f. Once the “operculum” (O) has detached from the capsule, the anterior chamber (A) prolapses outward, allowing the coiled tubule (T) to evert through a pore (arrow). g.—l. TEMs of undischarged nematocysts in intact cells. k. An identical striated banding pattern in the capsule wall in Polykrikos kofoidii (top), and Nematodinium sp. (bottom).   61	  62	Figure 4.5 Nematocysts in Nematodinium sp. a., d.,e., f. i.—l. TEMs in longitudinal (a., d.), oblique (f.), and transverse (e., g.-j.) section. b., c., g, h.: 3-D digital reconstructions from FIB-SEM on high-pressure frozen nematocysts. A = axial shaft. C = cap. P = plug. R = rosette. S = subcapsule. Scale bars = 1 µm.     Figure 4.6 Comparison of nematocyst characteristics in cnidarians and dinoflagellates. a. Extrusome features. b. Developmental features, with hydrozoan diagram redrawn from Holstein 1981, myxozoan diagram redrawn from Lom and DePuytorac 1965, polykrikoid diagram redrawn from Westfall, Bradbury and Townsend 1983, and first and second stages of the Nematodinium diagram redrawn from Greuet 1977, with the third stage drawn from observations made in this study. c. Discharge mechanism. Orange = nematocyst thread; gray = nematocyst capsule; blue = host cytoplasm.  63	  4.4 Discussion  4.4.1 Independent origins of pressurized ballistics.  While it is impossible to completely disprove a symbiogenic origin for nematocysts without sequencing every dinoflagellate genome, our findings have falsified the hypotheses made by Shostak (1993) and Hwang (2008) and favor independent origins for the nematocysts in dinoflagellates and cnidarians. In particular, Shostak hypothesized that cnidarians gained nematogenic genes by lateral gene transfer from endosymbiotic dinoflagellates (specifically Symbiodinium), but we found no known nematogenic genes in the genome of Symbiodinium sp. Minicollagens and other known nematogenic genes were also unrepresented among the genomes and transcriptomes available from other protists (Figure 4.3).   In a similar vein, we found no ultrastructural evidence of homology between cnidarian nematocysts and other ballistic structures that have previously been construed as cnidarian homologs (such as microsporidian polar filaments, which were once erroneously grouped with cnidarians in the Cnidosporia). Critically, poly-gamma-glutamate synthase (the enzyme that synthesizes osmotic propellant for nematocysts in cnidarians) was absent in the genomes of other organisms with osmotically-pressurized ballistics (e.g., microsporidians and helicosporidians). Therefore, pressurized ballistics appear to have been converged upon multiple times, both at the level of entire pressurized cells (microsporidians and helicosporidians) and organelles (nematocysts from dinoflagellates and ciliates).      64	4.4.2 Complexity through duplication  While they are fundamentally different from those of cnidarians, dinoflagellate nematocysts also differ vastly among dinoflagellate taxa. For instance, those of Nematodinium sp. have radial symmetry with manifold substructures that show no obvious homology to other dinoflagellate extrusomes (Figure 4.5). Based on these seemingly irreconcilable differences, Greuet and Hovasse (1977) suggested that nematocysts evolved multiple times within dinoflagellates. Since then, a new dinoflagellate extrusome has been described with multiple subunits—the docidosome of Actiniscus pentasterias, which bears two parallel internal ribbons (Hansen 1993). This fact, coupled with our observation that the nematocysts of Nematodinium sp. can have between seven (Greuet 1977) and fourteen internal subunits (Figure 4.5i-l), suggests that it is developmentally possible for dinoflagellates to multiply these substructures, and therefore attain greater extrusome complexity and size.   We hypothesize that the nematocysts of polykrikoids and Nematodinium sp. descended from an ancestral extrusome with only one inner capsule, which later underwent several multiplications in the ancestors of Nematodinum sp. There is already some evidence for homology between polykrikoid and warnowiid nematocysts (summarized in figure 4.6), and we add the strongest evidence of homology yet: a striated capsule wall composition that is identical in both Polykrikos kofoidii and Nematodinium sp. (Figure 4.4k).    4.4.3 Concluding remarks It seems that harpoon-like ballistics are a recurring theme in cellular evolution, with the most elaborate extrusomes evolving not in animals, but in predatory microbes. While the benefits of microscopic ballistics are not obvious (except in parasites that infect their hosts through explosive sporulation), there is evidently a strong pressure to evolve these structures—as they can arise through both autogenous routes, or even from specialized episymbionts (bacteria that can  65	extrude their contents as a projectile stream of cytoplasm, potentially to deter predators; Breglia, Yubuki, Hoppenrath and Leander 2010).   While defensive roles have been shown for a few extrusive structures (Rosati et al. 1999) and hypothesized for many more (Hausmann 1978), extrusomes can have other roles—as illustrated in predatory ciliates and polykrikoids (Matsuoka, Thomas, and Jacobson 2000; Morelli, Ricci and Verni 2002). Across eukaryotic diversity, most extrusome-bearing taxa have only one extrusome type per cell (Hausmann 1978), but polykrikoids have four (nematocysts, taeniocysts, mucocysts and trichocysts) and Nematodinium sp. has three (all but taeniocysts). Because both these groups are predators of other extrusome-bearing dinoflagellates (Gavelis et al. 2015b), perhaps an arms race is occurring between dinoflagellate predators and dinoflagellate prey. In other words, defensive features such as trichocysts and mucocysts may have prompted dinoflagellate predators to evolve more rapid extrusome types. These could help to circumvent prey defenses. Such an arms race seems to have occurred in predatory ciliates, in which taxa with the greatest extrusome diversity (e.g., Didinium, with three extrusome types, Hausmann 1978) prey on other well-armed ciliates. In a successful predation encounter, Didinium discharges its toxicysts before its prey is able to respond with trichocysts or mucocysts. Toxicysts deliver chemicals that disrupt the electrochemical signaling that the prey cell needs to coordinate its defenses. Perhaps the taeniocysts in polykrikoids function like toxicysts in Didinium, and incapacitate prey. The harpoonlike nematocyst seems to be used to reel the prey into the cytostome (Matsuoka 2001). This tethering feature could be important for capturing fast prey, since they might otherwise swim away before paralytic chemicals could take effect. Clearly more live observations of hunting behavior are needed.  While it is too soon to say what drives the evolution of this remarkable cellular armature, our findings show that a horizontal organellar transfer of nematocysts from cnidarians to dinoflagellates is much less likely than has historically been  66	assumed and suggests a route by which extreme subcellular complexity can be generated autogenously in a stepwise manner.                                 67	5 Conclusion  5.1 Contributions to understanding the evolution of complex organelles in dinoflagellates.  The work herein sheds light on the evolutionary origins of several eukaryotic organelles: the aberrant plastids of Polykrikos lebouriae, complex extrusomes of Polykrikos kofoidii and Nematodinium sp., and the eye-like ocelloids of Nematodinium sp., Warnowia sp., and Erythropsidinium agile. We combined different microscopic methods (including light microscopy, SEM, TEM, and FIB-SEM) with novel approaches using single organelle genomics and single cell transcriptomics.  Prior to this study, only PCR and TEM had been employed on these systems. In isolation, these methods could not determine whether nematocysts and ocelloids of dinoflagellates were homologous to the nematocysts and camera-type eyes of animals, respectively, or if they were the result of evolutionary convergence (Leander 2008b). Through novel microscopic and molecular approaches, we demonstrated the very different evolutionary trajectories from which these subcellular novelties have arisen in polykrikoids and warnowiids. As a means to this end, we also implemented the first single-cell use of spliced leader-primed PCR, as well as dissected organelle genomics.  One of the most significant discoveries was the plastid origin of the retinal body in Nematodinium sp., as described in Chapter 3. This was especially interesting given its close relationship to a mitochondrial “corneal” layer, which together show that the ocelloid incorporates elements from across multiple endosymbiotic origins. Yet these plastids and mitochondria are by no means new, given that all mitochondria stem from the last (or possibly first) eukaryotic common ancestor (Koonin 2015), and this particular plastid lineage dates at least as far back as the most recent common ancestor of dinoflagellates and apicomplexans  68	(Janouskovec et al. 2015). In other words, the eye-like nature of the ocelloid lies not its component organelles, but in the way they are arranged. Given the absence of structural genes (e.g., for proteins such as actin, tubulin or FtsZ) in the peridinin plastid genome, the shape of dinoflagellate plastids is clearly imposed extrinsically by nucleus-encoded proteins. A dizzying array of peridinin plastid morphologies have been generated in this way, with previous authors interpreting the plastids of Polykrikos lebouriae as kleptoplastids from diatoms or haptophytes (as described in Chapter 2) or confusing the plastid-derived retinal body of warnowiids with the eye of a jellyfish (described in Kofoid and Swezy 1921).   Some of our findings in this area were confirmatory of earlier discoveries in warnowiids. For instance, my transmission electron micrographs of Nematodinium sp. verified earlier data published by Greuet, who depicted the retinal body’s waveform membranes and its division by binary fission. However, we significantly advanced this understanding by extending our structural descriptions into three dimensions through FIB-SEM. We showed that the corneal mitochondria are not typical mitochondria (as stated by Greuet), but in fact form a continuous, laminar layer surrounding the lens, as essentially a single “corneal” mitochondrion. Likewise, we found that the retinal body was in fact continuous with numerous plastids throughout the cytoplasm. This discovery of a “plastid network” greatly raises our appreciation of structural complexity in organelles, for its continuity with the retinal body makes the plastid network—to our knowledge—the single most complex plastid described. Thus, the ocelloid defines a new extreme of plastid differentiation, and shows the great malleability of peridinin plastids.   Lastly, this chapter included significant methodological novelty, as we employed MDA on dissected organelles, rather than single cells, for which MDA was intended. This allowed us to carry out an experimental approach rarely possible in uncultivated cells. In doing so, we procured the first genomic data from  69	warnowiid dinoflagellates, and addressed longstanding questions about ocelloid evolution.  In Chapter 4, our investigation of nematocysts in polykrikoids and Nematodinium sp. suggested that the evolution of complex extrusomes traversed a different path, by multiplying existing features rather than using symbiotically acquired components. We found that the structure and function of nematocysts in Polykrikos kofoidii is both fundamentally different and more complex than in cnidarians, and shows clear homology to the nematocysts of other dinoflagellates, such as Nematodinium sp. This entailed the first SEM study of dinoflagellate nematocysts: including in their discharged states. These data demonstrated the absence of a stylet and hatchlike operculum, contrary to previous descriptions by Westfall (1985). While Westfall stated that these structures appeared similar to stylets and opercula in cnidarian, our SEM analysis of nematocyst discharge showed that they lack an obvious shared function, as the ‘operculum’ does not open in a hatchlike way (rather, the thread exits through a central pore), and the stylet does not extrude outside the capsule to deliver toxins envenomation.   Our initial TEM data in Nematodinium sp. and P. kofoidii largely confirmed findings by Greuet (1973) and Westfall (1985), though we found some important new features. This included the striated pattern in the nematocyst capsule of P. kofoidii that, in our view, constitutes an unambiguous synapomorphy for nematocysts in the polykrikoid-warnowiid clade. This constitutes the strongest evidence of nematocyst monophyly within dinoflagellates. Through FIB-SEM, we also produced the first description of a fully mature nematocyst in Nematodinium sp. This structure possessed a differentiated apical region (diagramed in Fig. 4.5) that had eluded description by previous authors. In sum, our structural findings underscored the fundamental differences between nematocysts in dinoflagellates and cnidarians.   70	 In addition to morphological work, we exhaustively searched available dinoflagellate transcriptomes and several representative eukaryote genomes for cnidarian nematocysts proteins, in order to test for possible homology between the nematocysts of dinoflagellates and cnidarians. The absence of such proteins in non-animal genomes strongly supports the alternative hypothesis: that nematocysts evolved independently in dinoflagellates and cnidarians. Our analysis supports the prevalence of convergent evolution in eukaryotic history, which has evidently allowed for multiple origins of advanced ballistic structures in microbial predators and parasites.   Lastly, we considered how the multi-capsule nematocyst of Nematodinium sp. could have originated from a simpler precursor through the duplication of internal components (Figure 4.5). The piecemeal assembly of these nematocysts in endomembrane vesicles strongly testifies to their autogenous origin, and exhibits a level of vesicular coordination that has not been observed in other systems. Together, the ocelloid and nematocysts define extremes in the evolution of subcellular complexity—one through endosymbiosis, and the other, evidently, by autogenous processes.  5.2 Future directions   Throughout these studies, one major obstacle was the preferential amplification of bacterial genomic “contaminants” by MDA. This was a drawback when searching for plastid genomes (as in Chapters 2 and 3), but will prove to be an advantage for anyone studying bacteria from the wild. In keeping with this, I have begun to characterize the bacterial symbionts of the spectacular tropical dinoflagellates Ornthicercus, Amphisolenia, and Podolampas, which—with the help of their symbionts—perform essential nitrogen-fixing roles in their oligotrophic habitats, and may serve other roles that have yet to be uncovered (such as photosynthesis, indicated by attenuated thylakoid-like membranes).    71	Another recent opportunity has been the establishment of Polykrikos kofoidii in culture by Tillmann and Hoppenrath (2013). The newfound cultivability of polykrikoids may allow their nematocysts to be harvested in sufficient quantities for proteomic analysis, in order to reveal the constituents of these fantastically complex structures. A genetic transformation system for polykrikoids would also be highly desirable, because so much has been learned about the extrusomes of ciliates through careful knockout studies of proteins involved in extrusome biogenesis (Chilcoat et al. 1996.) Of course, higher phylogenetic resolution will also be needed for reconstructions of dinoflagellate character evolution, e.g., if we are to understand the sequence of events that produced the warnowiid ocelloid (Figure 5.1).  Figure 5.1. Dinoflagellate eyespot types within a phylogenetic context. Diagrams of whole cells and eyespots are shown for all dinoflagellates for which both ultrastructural descriptions and 18S and 28S ribosomal rDNA sequences have been published. Maximum likelihood and Bayesian support values above 70/.7 are shown. Eyespot diagrams highlight plastid-like structures (crimson), as well as mitochondria (dark blue), lens-like vesicles (light blue), lipid droplets (red dots), and crystalline layers (grey dashes).   72	Live observation will be essential to understand the uses of these complex organelles. Very little is known about the prey capture mechanism of polykrikoids. Our understanding of this process could be clarified by recording prey capture using DIC light microscopy (and a higher resolution than in Hwang et al. [2008] and Matsuoka et al. [2001]), as well as high-speed microvideography. Most importantly, behavioral experiments are needed to confirm our hypothesis that the ocelloid is involved in polarized light detection, and this could be tested either through observations of captured warnowiids (specifically with tropical species that can survive at room temperature), or field-based studies, e.g., through the development—but first, the invention—of polarized light traps. Ultimately, there is no shortage of interesting questions to ask about dinoflagellates, only a shortage of proven techniques for studying them.      73	Bibliography   Adl, S. M., Leander, B.S., Simpson, A.G.B., Archibald, J.M., Anderson, O.R., Bass, D., Bowser, S.S., et al. 2007. Diversity, nomenclature, and taxonomy of protists. Syst. Biol., 56:684-689.  Bachvaroff, T. R., Sanchez-Puerta, M. V., and Delwiche, C. F. 2006. Rate variation as a function of gene origin in plastid-derived genes of peridinin-containing dinoflagellates. Journal of molecular evolution. 62(1): p. 42-52.  Boisvert, S., Laviolette, F., and Corbeil, J. 2010. Ray: Simultaneous assembly of reads from a mix of high-throughput sequencing technologies. Journal of Computational Biology. 17(11): p. 1519-1533.  Bourlat, S. J., Nielsen, C., Lockyer, A. E., Littlewood, D. T. J., and Telford, M. J. 2003. Xenoturbella is a deuterostome that eats molluscs. Nature. 424(6951): p. 925-928.  Breglia, S. A., Yubuki, N., Hoppenrath, M., and Leander, B. S. (2010). Ultrastructure and molecular phylogenetic position of a novel euglenozoan with extrusive episymbiotic bacteria: Bihospites bacati n. gen. et sp.(Symbiontida). BMC Microbiology. 10(1): p. 145.  Brendtsen, J.D., Nielsen, H., von Heijne, G., and Brunak, S. 2004. Improved prediction of signal peptides: SignalP 3.0. Journal of Molecular Biology. 340(4): p. 783-95.  Burki, F. 2014. The eukaryotic tree of life from a global phylogenomic perspective. Cold Spring Harbor perspectives in biology. 6(5): p. a016147.  Castresana, J. 2000. Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Molecular Biology and Evolution. 17(4): p. 540-552.  Chilcoat, N. D., Melia, S. M., Haddad, A., and Turkewitz, A. P. 1996. Granule lattice protein 1 (Grl1p), an acidic, calcium-binding protein in Tetrahymena thermophila dense-core secretory granules, influences granule size, shape, content organization, and release but not protein sorting or condensation. The Journal of Cell Biology. 135(6): p. 1775-1787.  Corradi, N., Akiyoshi, D. E., Morrison, H. G., Fend, X., Weiss, L. M., Tzipori, S., and Keeling, P. J. 2007. Patterns of genome evolution between the compacter genomes of the microsporidian parasites Encephalitozoon cuniculi, Antonospora locustae and Enterocytozoon bieneusi. PLoS One. 2(12): p. e1277.   74	Dacks, J. B., and Field, M. C. 2007. Evolution of the eukaryotic membrane-trafficking system: origin, tempo and mode. Journal of cell science. 120(17): p. 2977-2985.  David, C. N., Özbek, S., Adamczyk, P., Meier, S., Pauly, B., Chapman, J., ... and Holstein, T. W. 2008. Evolution of complex structures: minicollagens shape the cnidarian nematocyst. Trends in Genetics. 24(9): p. 431-438.  Dodge, J. D. 1984. The functional and phylogenetic significance of dinoflagellate eyespots. Biosystems. 16(3): p. 259-267.  Edgar, R.C. 2004. MUSCLE: a multiple sequence alignment method with reduced time and space complexity. BMC Bioinformatics. 5: p. 1-19.  Fensome, R. A., Norris, G., Sarjeant, W. A. S., Taylor, F. J. R., Wharton, D. I., and Williams, G. L. 1993. A classification of Living and Fossil Dinoflagellates. Micropaleontology Press, American Museum of Natural History.  Foster, K. W., and Smyth, R. D. 1980. Light antennas in phototactic algae. Microbiological Reviews. 44(4): p. 572-630.  Gavelis, G. S., White, R. A., Suttle, C. A., Keeling, P. J., and Leander, B. S. 2015. Single-cell transcriptomics using spliced leader PCR: Evidence for multiple losses of photosynthesis in polykrikoid dinoflagellates. BMC Genomics.16(1): p. 528.  Gomez, F., Lopez-Garcia, P., and Moreira, D. 2009. Molecular phylogeny of the ocelloid‐bearing dinoflagellates Erythropsidinium and Warnowia (Warnowiaceae, Dinophyceae). Journal of Eukaryotic Microbiology. 56(5): p. 440-445.  Gornik, S. G., Ford, K. L., Mulhern, T. D., Bacic, A., McFadden, G. I., and Waller, R. F. 2012. Loss of nucleosomal DNA condensation coincides with appearance of a novel nuclear protein in dinoflagellates. Current Biology. 22(24): p. 2303-2312.  Gehring, W. J. 2005. New perspectives on eye development and the evolution of eyes and photoreceptors. Journal of Heredity. 96(3): p. 171-184.  Greuet, C., and Hovasse, R. 1977. About genesis of nematocysts of Polykrikos schwartzii. Protistologica. 13(1): p. 145-149.  Greuet, C. 1971. Ultrastructural study and evolution of cnidocysts Nematodinium, péridinien Warnowiidae Lindemann. Protistologica. 7(3): p. 345-355.  Greuet, C. 1977. Structural and ultrastructural evolution of ocelloid of Erythropsidinium-pavillardi-Kofoid-and-Swezy (dinoflagellate Warnowiidae,  75	Lindemann) during division and palintomic divisions. Protistologica. 13(1): p. 127-143.  Greuet, C. 1987. Dinoflagellate ultrastructure and complex organelles. In The Biology of Dinoflagellates (Dodge, J., ed.), Blackwell Scientific Publications pp. 92-142.  Hansen, G. (1993). Light and electron microscopical observations of the dinoflagellate Actiniscus pentasterias (Dinophyceae). Journal of Phycology. 29(4): p. 486-499.  Hausmann, K. (1978). Extrusive Organelles in Protists. International Review of Cytology. 52: p. 197-269.  Holstein, T. 1981. The morphogenesis of nematocytes in Hydra and Forsklia: An ultrastructural study. Journal of Ultrastructure Research. 75(3): p. 276-290.  Hoppenrath, M., et al., 2010. Re-classification of Pheopolykrikos hartmannii as  Polykrikos (Dinophyceae) based partly on the ultrastructure of complex  extrusomes. European Journal of Protistology. 46(1): p. 29-37.  Hoppenrath, M., Bachvaroff, T. R., Handy, S. M., Delwiche, C. F., and Leander, B. S. (2009). Molecular phylogeny of ocelloid-bearing dinoflagellates (Warnowiaceae) as inferred from SSU and LSU rDNA sequences. BMC evolutionary biology.9(1): p. 116.  Hoppenrath, M. and Leander, B.S. 2007a. Morphology and phylogeny of the  pseudocolonial dinoflagellates Polykrikos lebouriae and Polykrikos herdmanae n.  sp. Protist. 158(2): p. 209-227.  Hoppenrath, M. and Leander B.S. 2007b. Character evolution in polykrikoid  dinoflagellates. Journal of Phycology. 43(2): p. 366-377.  Hovasse, R. 1951. Contribution à l’étude de la cnidogénèse chez les péridiniens. I. Cnidogénèse cyclique chez Polykrikos schwartzi. Archives de Expérimental et General Zoologie. 87: p. 299.  Howe, C. J., Nisbet, R. E. R., and Barbrook, A. C. 2008. The remarkable chloroplast genome of dinoflagellates. Journal of experimental botany. 59(5): p. 1035-1045.  Huelsenbeck, J.P. and Ronquist, F. 2001. MRBAYES: Bayesian inference of phylogenetic trees. Bioinformatics. 17(8): p. 754-755.  76	 Hwang, J. S., Nagai, S., Hayakawa, S., Takaku, Y., and Gojobori, T. 2008. The search for the origin of cnidarian nematocysts in dinoflagellates. In Evolutionary Biology from Concept to Application (p. 135-152). Springer Berlin Heidelberg.  Imanian, B., Pombert, J. F., and Keeling, P. J. 2010. The complete plastid genomes of the two ‘dinotoms’ Durinskia baltica and Kryptoperidinium foliaceum. PloS one. 5(5): p. e10711.  Janouškovec, J., Horák, A., Oborník, M., Lukeš, J., and Keeling, P. J. 2010. A common red algal origin of the apicomplexan, dinoflagellate, and heterokont plastids. Proceedings of the National Academy of Sciences. 107(24): p. 10949-10954  Janouškovec, J., Horák, A., Barott, K. L., Rohwer, F. L., and Keeling, P. J. 2013. Environmental distribution of coral-associated relatives of apicomplexan parasites. The ISME journal, 7(2): p. 444-447.  Janouškovec, J., Tikhonenkov, D. V., Burki, F., Howe, A. T., Kolísko, M., Mylnikov, A. P., and Keeling, P. J. 2015. Factors mediating plastid dependency and the origins of parasitism in apicomplexans and their close relatives.Proceedings of the National Academy of Sciences. p. 201423790.  Kang, N. S., Jeong, H. J., Moestrup, Ø., Shin, W., Nam, S. W., Park, J. Y., ... and Noh, J. H. 2010. Description of a new planktonic mixotrophic dinoflagellate Paragymnodinium shiwhaense n. gen., n. sp. from the coastal waters off western Korea: morphology, pigments, and ribosomal DNA gene sequence. Journal of Eukaryotic Microbiology. 57(2): p. 121-144.  Kang, N. S., Jeong, H. J., Moestrup, Ø., and Park, T. G. 2011. Gyrodiniellum shiwhaense n. gen., n. sp., a new planktonic heterotrophic dinoflagellate from the coastal waters of western Korea: morphology and ribosomal DNA gene sequence. Journal of Eukaryotic Microbiology, 58(4), 284-309.  Keeling, P. J., Leander, B. S., and Lukeš, J. 2010. Reply to Speijer: Does complexity necessarily arise from selective advantage? Proceedings of the National Academy of Sciences. 107(7): p. E26.  Keeling, P. J. 2010. The endosymbiotic origin, diversification and fate of plastids. Philosophical Transactions of the Royal Society of London B: Biological Sciences. 365(1541), 729-748.  Kofoid, C. A., and Swezy, O. 1921. The free-living unarmored Dinoflagellata. Regents of the University of California, Berkeley.    77	Kolisko, M., Boscaro, V., Burki, F., Lynn, D. H., and Keeling, P. J. 2014. Single-cell transcriptomics for microbial eukaryotes. Current Biology. 24(22): p. R1081-R1082.  Koonin, E. V. 2015. Origin of eukaryotes from within archaea, archaeal eukaryome and bursts of gene gain: eukaryogenesis just made easier?. Philosophical Transactions of the Royal Society Series B. 370(1678): 20140333.  Langmead, B., Trapnell, C., Pop, M., and Salzberg, S. L. 2009. Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biology. 10(3): R25.  Leander, B. S., Saldarriaga, J. F., and Keeling, P. J. 2002. Surface morphology of the marine parasite Haplozoon axiothellae Siebert (Dinoflagellata). European Journal of Protistology. 38(3): p. 287-297.  Leander, B. S. 2008. A hierarchical view of convergent evolution in microbial eukaryotes. Journal of Eukaryotic Microbiology. 55(2): p. 59-68.  Leander, B. S. 2008. Different modes of convergent evolution reflect phylogenetic distances: a reply to Arendt and Reznick. Update. 23(9).  Liu, J., and Kattawar, G. W. 2013. Detection of dinoflagellates by the light scattering properties of the chiral structure of their chromosomes. Journal of Quantitative Spectroscopy and Radiative Transfer. 131: p. 24-33.  Lom, J., and dePuytorac, P. 1965. Studies on the myxosporidian ultrastructure and polar capsule development. Protistologica. 1(1): p. 53-65.  Mast, F. D., Barlow, L. D., Rachubinski, R. A., and Dacks, J. B. 2014. Evolutionary mechanisms for establishing eukaryotic cellular complexity. Trends in cell biology. 24(7): p. 435-442.  Matsuoka, K., Cho, H. J., and Jacobson, D. M. 2000. Observations of the feeding behavior and growth rates of the heterotrophic dinoflagellate Polykrikos kofoidii (Polykrikaceae, Dinophyceae). Phycologia, 39(1). p. 82-86.  Meyer, F., et al. 2004. The metagenomics RAST server - a public resource for the automatic phylogenetic and functional analysis of metagenomes. BMC Bioinformatics. 2008. 9.  Minge, M. A., Shalchian-Tabrizi, K., Tørresen, O. K., Takishita, K., Probert, I.,  Inagaki, Y., ... and Jakobsen, K. S. 2010. A phylogenetic mosaic plastid proteome  and unusual plastid-targeting signals in the green-colored dinoflagellate  Lepidodinium chlorophorum. BMC evolutionary biology, 10(1): p. 191.  78	 Mornin, L., and Francis, D. (1967). Fine struction of Nematodinium armatum, a naked dinoflagellates. Journal of Microscopy. 6(6): p. 759-772.  Nassoury, N., Cappadocia, M., and Morse, D. 2003. Plastid ultrastructure defines the protein import pathway in dinoflagellates. Journal of Cell Science. 116(14): p. 2867-74.  Nielsen, H. and Krogh, A. 1998. Prediction of signal peptides and signal anchors by a hidden Markov model. International Conference on Intelligent Systems for Molecular Biology. 6: p. 122-30.  Patron, N.J., Waller, R.F., Archibald, J.M, and Keeling, P.J. 2005. Complex protein targeting to dinoflagellate plastids. Journal of Molecular Biology. 348(4): p. 1015-24.  Qiu, D., Huang, L., Liu, S., Zhang, H., and Lin, S. 2013. Apical Groove Type and  Molecular Phylogeny Suggests Reclassification of Cochlodinium geminatum as  Polykrikos geminatum. Plos One. 8(8).  Morelli, A., Ricci, N., and Verni, F. 2002. Action of Litonotus (predator) toxicysts on the electric properties of Euplotes (prey) cell membrane. Italian Journal of Zoology. 69(2): 103-107.  Raikov, I. B. (1992). Unusual extrusive organelles in karyorelictid ciliates: an argument for the ancient origin of this group. BioSystems. 28(1): p. 195-201.  Ronquist, F. and Huelsenbeck, J.P. 2003. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics. 19(12): p. 1572-1574.  Rosati, G., Petroni, G., Quochi, S., Modeo, L., and Verni, F. 1999. Epixenosomes: peculiar epibionts of the hypotrich ciliate Euplotidium itoi defend their host against predators. Journal of Eukaryotic Microbiology. 46(3): 278-282.  Saldarriaga, J.F., Taylor, F.J.R., Keeling, P.J., and Cavalier-Smith, T. 2001. Dinoflagellate nuclear SSU rRNA phylogeny suggests multiple plastid losses and replacements. Journal of Molecular Evolution. 53(3): p. 204-213.  Schrallhammer, M., Schweikert, M., Vallesi, A., Verni, F., and Giulio P. 2011. Detection of a novel subspecies of Francisella noatunensis as endosymbiont of the ciliate Euplotes raikovi. Microbial Ecology, 2011. 61(2): p. 455-464.  Shoguchi, E., Shinzato, C., Kawashima, T., Gyoja, F., Mungpakdee, S., Koyanagi, R., ... and Satoh, N. 2013. Draft assembly of the Symbiodinium minutum nuclear genome reveals dinoflagellate gene structure. Current Biology. 23(15): p. 1399-1408.  79	 Shostak, S. 1993. A symbiogenetic theory for the origins of cnidocysts in Cnidaria. BioSystems. 29(1): p. 49-58.  Sonnhammer, E.L., von Heijne, G., and A. Krogh. 1998. A hidden Markov model for predicted transmembrane helices in protein sequences. International Conference on Intelligent Systems for Molecular Biology. (6): p. 175-82.  Stamatakis, A. 2006. RAxML-VI-HPC: Maximum likelihood-based phylogenetic analyses with thousands of taxa and mixed models. Bioinformatics. 22(21): p. 2688-2690.  Tamura, K., Stecher, G., Peterson, D., Filipski, A., and Kumar, S. 2013. MEGA6: molecular evolutionary genetics analysis version 6.0. Molecular Biology and Evolution. 30(12): p. 2725-2729.  Tillmann, U., and Hoppenrath, M. 2013. Life cycle of the pseudocolonial dinoflagellate Polykrikos kofoidii (Gymnodiniales, Dinoflagellata). Journal of Phycology. 49(2): p. 298-317.  Uhlig, G., Thiel, H., and Gray, J.S. 1973. Quantitative separation of meiofauna - comparison of methods. Helgolander Wissenschaftliche Meeresuntersuchungen. 25(1): p. 173-195.  Vivier, E. and Desportes, I. 1990. Phylum Apicomplexa. In The Handbook of Protoctista (Margulis, L., ed.), Jones and Bartlett Publishers pp. 549-573.  Widder, E. A. 2010. Bioluminescence in the ocean: origins of biological, chemical, and ecological diversity. Science. 328(5979): p. 704-708.  Yamaguchi, A., Yubuki, N., and Leander, B. S. 2012. Morphostasis in a novel eukaryote illuminates the evolutionary transition from phagotrophy to phototrophy: description of Rapaza viridis n. gen. et sp. (Euglenozoa, Euglenida). BMC Evolutionary Biology. 12(1): p. 29.  Yoon, H. S., Price, D. C., Stepanauskas, R., Rajah, V. D., Sieracki, M. E., Wilson, W. H., ... and Bhattacharya, D. 2011. Single-cell genomics reveals organismal interactions in uncultivated marine protists. Science. 332(6030): p. 714-717.  Zhang, H., Zhuang, Y., Gill, J., and Lin, S. 2007. Spliced leader RNA trans-splicing in dinoflagellates. Proceedings of the National Academy of Sciences of the United States of America. 104(11): p. 4618-4623.  Zhang, H., et al. 2013. Proof that dinoflagellate spliced leader (dinoSL) is a useful hook for fishing dinoflagellate transcripts from mixed microbial samples:  80	Symbiodinium kawagutii as a case study. Protist. 164(4): p. 510-527.      


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