Ciliate Biodiversity and Phylogenetic Reconstruction Assessed by Multiple Molecular Markers Micah Dunthorn University of Massachusetts Amherst, Dunthorn@Nsm.Umass.Edu

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9-2009 Ciliate Biodiversity and Phylogenetic Reconstruction Assessed by Multiple Molecular Markers Micah Dunthorn University of Massachusetts Amherst, [email protected]

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CILIATE BIODIVERSITY AND PHYLOGENETIC RECONSTRUCTION

ASSESSED BY MULTIPLE MOLECULAR MARKERS

A Dissertation Presented

MICAH DUNTHORN

Submitted to the Graduate School of the University of Massachusetts Amherst in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

September 2009

Organismic and Evolutionary Biology

CILIATE BIODIVERSITY AND PHYLOGENETIC RECONSTRUCTION

ASSESSED BY MULTIPLE MOLECULAR MARKERS

A Dissertation Presented

MICAH DUNTHORN

Approved as to style and content by:

______Laura A. Katz, chair

______Robert L. Dorit, member

______George B. McManus, member

______Benjamin B. Normark, member

______Joseph S. Elkinton, Graduate Program Director Program in Organismic & Evolutionary Biology

To Frank Buchholz

13 February 1952—15 July 1991

ACKNOWLEDGEMENTS

Much thanks goes to my dissertation committee. Laura A. Katz, my advisor, provided her support throughout this dissertation. I thank Laura for allowing me to work in her lab and for teaching me many molecular methods. Rob Dorit was able to point out the bigger picture and context. George McManus was great to talk about ciliate evolution and the intricacies of ciliate taxonomy. And Ben Normark was always there to push me towards our current theoretical gaps.

Much thanks also goes to Professor Dr. Wilhelm Foissner, who isolated and identified most of the ciliates that were used in this dissertation, and is a co-author on many of the chapters. I am not sure how he was able to put up with me while I visited his laboratory. Chapter 2 was a collaboration with Thorsten Stoeck, Marion Eppinger, M. V.

Julian Schwarz, Michael Schweikert, and Jens Boenigk. Denis Lynn collaborated on the

PhyloCode classification in Chapter 7. Phil Cantino, the editor for the Companion

Volume to which the PhyloCode definitions will be published, was extremely helpful and patient in his edits.

This dissertation would also not be possible without the many folks in the Katz lab. I will miss Mary Doherty’s jokes and her insights into ciliate biogeography. The following directly or indirectly also made an impact: Becky Zufall, Sofia Annis, Erica

Barbero, Jennifer DeBarardinis, Jessica Grant, Meaghan Hall, Hannah Jaris, Allie

Kovner, Dan Lahr, Christina Lyman, Bob Merritt, Laura Parfrey, Maiko Tamura, Yonas

Tekle, Sam Torquato, and Laura Vann. Help also came from members of the Organismic and Evolutionary Biology program: Penny Jaques, Norman Johnson, David Lahti, and

v Lynn Margulis. Caffeine and delicious vegan snacks were made possible by Melissa

Krueger at the Elbow Room Café.

Partial funding for this dissertation and for presenting at conferences came from:

International Travel Award for Young Scholars, Society for the Study of Evolution;

Holz-Connor Travel Grant, International Society of Protozoologists; Mini-PEET Award,

Society of Systematic Biologists; Graduate Research Award, Society of Systematic

Biologists; Travel Grant, 56th Annual Meeting, Society of Protozoologists; Graduate

Fellowship, Massachusetts Space Grant Consortium. Funding was also provided by Laura

A. Katz through her U.S. National Science Foundation grant #636498.

I would also like to thank my mother and father—Lani and Tom—for their patience and help throughout my time in graduate school. Finally, I want to thank my son—(captain) Aureliano—for thinking that working in the lab consists of watching movies in my office, since that is what he did the so many times I dragged him to the lab so I can perform just more PCR and cloning on a Saturday afternoon.

vi ABSTRACT

CILIATE BIODIVERSITY AND PHYLOGENETIC RECONSTRUCTION

ASSESSED BY MULTIPLE MOLECULAR MARKERS

SEPTEMBER 2009

MICAH DUNTHORN, B.A., GEORGE MASON UNIVERSITY

M.A., UNIVERSITY OF MISSOURI ST. LOUIS

PH.D., UNIVERSITY OF MASSACHUSETTS AMHERST

Directed by: Laura A. Katz

Ciliates provide a powerful system within microbial eukaryotes in which molecular genealogies can be compared to detailed morphological taxonomies. Two groups with such detailed taxonomies are the Colpodea and the Halteriidae. There are about 200 described Colpodea species that are found primarily in terrestrial habitats. In

Chapters 1 and 2, taxon sampling is increased to include exemplars from all major subclades using nuclear small subunit rDNA (nSSU-rDNA) sequencing. Much of the morphological taxonomy is supported, but extensive non-monophyly is found throughout. The conflict between some nodes of the nSSU-rDNA genealogy and morphology-based taxonomy suggests the need for additional molecular marker. In

Chapter 3, character sampling is increased using mitochondrial small subunit rDNA

(mtSSU-rDNA) sequencing. The nSSU-rDNA and mtSSU-rDNA topologies for the

Colpodea are largely congruent for well-supported nodes, suggesting that nSSU-rDNA work in other ciliate clades will be supported by mtSSU-rDNA as well. Chapter 4 compares the underlying genetic variation within two closely related species in the

vii Halteriidae with increased taxon and molecular sampling using nSSU-rDNA and internally-transcribed spacer (ITS) region sequencing. The morphospecies Halteria grandinella shows extensive genetic variation that is consistent with either a large effective population size or the existence of multiple cryptic species. This pattern contrasts with the minimal of genetic variation in the morphospecies Meseres corlissi.

Chapter 5 discusses the congruence and incongruence among morphological and molecular data in ciliates. Most of the incongruence occurs where there is little statistical support for the molecules, or where molecular data is consistent with alternative morphological hypotheses. Chapter 6 reviews the data for sex, or lack thereof, in the

Colpodea, a potentially ancient asexual group where sex was regained in a derived species. In Chapter 7, four ciliate clades are redefined using the PhyloCode.

viii CONTENTS Page ACKNOWLEDGEMENTS……………………………………………………………… v

ABSTRACT…………………………………………………………………………..…vii

LIST OF TABLES………………………………………………………………………xiv

LIST OF FIGURES……………………………………………………………………...xv

CHAPTERS

1. MOLECULAR PHYLOGENETIC ANALYSIS OF CLASS COLPODEA (PHYLUM CILIOPHORA) USING BROAD TAXON SAMPLING………………….……………..1

1.1. Abstract……………………………………………………………………….2 1.2. Introduction……………………….…………………………………………..3 1.3. Materials and methods…………………….………………………...... 7

1.3.1. Taxon sampling and collection……………………………..………7 1.3.2. Identification………………………………………………………..7 1.3.3. DNA extraction, amplification, cloning and sequencing…………...7 1.3.4. Genealogical analyses………………………………………………9

1.4. Results……………………………………………………………………….10

1.4.1. Pairwise distances within collections……………………………...10 1.4.2. Deletion within one SSU rDNA copy in Bryometopus pseudochilodon…………………………………………………..11 1.4.3. Intron in Cyrtolophosis mucicola…………………………………12 1.4.4. SSU rDNA genealogy of the Colpodea…………………………...12

1.5. Discussion…………………………………………………………………...16

1.5.1. Comparisons between morphology and molecules………………..16

1.5.1.1. The class…………………………………………………16 1.5.1.2. The orders……………………………………………….16 1.5.1.3. The genus Colpoda……………………………………...20

1.5.2. Open questions with some species designations………………..…21 1.5.3. Evidence for sex…………………………………………………...22 1.5.4. Group I intron in Cyrtolophosis mucicola………………………...23 1.5.5. Reconciling morphology and molecules in the Colpodea………...24

1.6. References………………………………………………………………...…25

ix 2. PHYLOGENETIC PLACEMENT OF THE CYRTOLOPHOSIDIDAE STOKES, 1888 (CILIOPHORA; COLPODEA) AND NEOTYPIFICATION OF ARISTEROSTOMA MARINUM KAHL, 1931…………………………………………...40

2.1. Abstract……...………………………………………………………………41 2.2. Introduction………………………………………………………………….42 2.3. Methods……………………………………………………………………...43

2.3.1. Taxon sampling……………………………………………………43 2.3.2. Light and electron microscopy…………………………………….44 2.3.3. Terminology……………………………………………………….45 2.3.4. Autecology……………………………………………………...…45 2.3.5. Amplification and sequencing ……………………………………47 2.3.6. Genealogical analyses……………………………………………..48 2.3.7. Constrained analysis……………………………………………....49 2.3.8. Rate class analysis…………………………………………………49

2.4. Results……………………………………………………………………….49

2.4.1. Description of the neotype of Aristerostoma marinum Kahl, 1931………………………………………………………..49 2.4.2. Electron microscopy………………………………………………51 2.4.3. Autecology………………………………………………………..52 2.4.4. Pairwise SSU rDNA sequence differences within the Cyrtolophosididae………………………………………………..52 2.4.5. Genealogical analysis……………………………………………...53 2.4.6. Comparisons of hypotheses……………………………………….54 2.4.7. Long-branch attraction…………………………………………….54

2.5. Discussion…..……………………………………………………………….56

2.5.1. Neotypification and emended diagnosis of Aristerostoma marinum Kahl 1931………………………....…………………...55 2.5.2. Emended diagnosis………………………………………………..56 2.5.3. Occurrence and ecology……………………………………….…..57 2.5.4. Comparison with original descriptions and related species……….58 2.5.5. Neotype specimens…………………………………………..……61 2.5.6. Pairwise distance between Aristerostoma morphospecies………...61 2.5.7. Phylogenetic placement of the order Cyrtolophosidida…………...61 2.5.8. Reconciling morphology and molecules…………………………..62 2.5.9. Assignment of GenBank environmental SSU-rDNA sequences to the Cyrtolophosididae…………………………...…64

2.6. Conclusions………………………………………………………………….65 2.7. References…………………………………………………………………...67

x 3. EXPANDING CHARACTER SAMPLING IN CILIATE PHYLOGENETIC RECONSTRUCTION: MITOCHONDRIAL SSU-rDNA AS A MOLECULAR MARKER………………………………………………………………………………..86

3.1. Abstract……...………………………………………………………………87 3.2. Introduction………………………………………………………………….88 3.3. Methods……………………………………………………………………...90

3.3.1. Taxon sampling and terminology…………………………………90 3.3.2. DNA amplification and sequencing……………………………….91 3.3.3. Genealogical analyses……………………………………………..91 3.3.4. Data partitioning and congruence testing…………………………92

3.4. Results……………………………………………………………………….93

3.4.1. Characteristics of gene sequences…………………………………93 3.4.2. MtSSU-rDNA analyses……………………………………………94 3.4.3. Nuclear SSU-rDNA analyses and topology congruence………….96 3.4.4. Concatenated analyses…………………………………………….97

3.5. Discussion……………………………………………………………….…..98

3.5.1. Mitochondrial SSU-rDNA as a ciliate molecular marker…………98 3.5.2. Morphology vs. molecules in ciliates……………………………100 3.5.3. Systematic implications………………………………………….101

3.6. Conclusions………………………………………………………………...101 3.7. References………………………………………………………………….102

4. EXTENSIVE GENETIC DIVERSITY WITHIN HALTERIID CILIATES………...115

4.1. Abstract……...……………………………………………………………..116 4.2. Introduction………………………………………………………………...117 4.3. Methods…………………………………………………………………….118

4.3.1. Taxon sampling and identification……………………………….118 4.3.2. Amplification, cloning, and sequencing…………………………119 4.3.3. Genealogical analyses……………………………………………120

4.4. Results……………………………………………………………………...121

4.4.1. Intra-isolate pairwise distances…………………………………..121 4.4.2. Genealogies………………………………………………………122

4.5. Discussion………………………………………………………………….124

4.4.1. Genetic variation underlying morphospecies…………………….124

4.6. References………………………………………………………………….126

5. RICHNESS OF MORPHOLOGICAL HYPOTHESES IN CILIATE SYSTEMATICS ALLOWS FOR DETAILED ASSESSMENT OF HOMOLOGY AND COMPARISONS WITH GENE TREES………………………………………...138

5.1. Abstract……...……………………………………………………………..139 5.2. Introduction………………………………………………………………...140 5.3. Strengths of morphology…………………………………………………...140

5.3.1. Species delimitations…………………………………………….140 5.3.2. Ciliate tree of life………………………………………………...141

5.4. Challenges to Morphological analyses…………………………………….142

5.4.1. Decline in number of trained taxonomists……………………….142 5.4.2. Number of characters…………………………………………….142 5.4.3. Homology assessment……………………………………………143

5.5. Concordance with molecular hypotheses …………………………………..144 5.6. Open Questions…………………………………………………………….145

5.6.1. Lack of morphological variation when there is genetic diversity..145 5.6.2. Lack of genetic diversity when there is morphological variation..146

5.7. References………………………………………………………………….148

6. ANCIENT ASEXUAL LINEAGE OF CILIATES THAT REGAINED SEX?...... 157

6.1. Abstract……...……………………………………………………………..158 6.2. Introduction………………………………………………………………...159 6.3. Empirical evidence…………………………………………………………160 6.4. Are colpodeans ancient asexuals? …………………………………………162 6.5. Did colpodeans reverse the loss of sex? …………………………………..163 6.6. Conclusion…………………………………………………………………165 6.7. References………………………………………………………………….166

xii 7. PHYLOCODE DEFINITIONS FOR FOUR CILIATE CLADES..…………………173

7.1. Abstract……...………………………………………………………….….174 7.2. Introduction………………………………………………………………...175 7.3. The PhyloCode……………………………………………………………..176 7.4. Why the PhyloCode should be applied to ciliate…………………………..177 7.5. Application of the PhyloCode to four ciliate clades……………………….179

7.5.1. Ciliophora………………………………………………………..180 7.5.2. Postciliodesmatophora…………………………………………..182 7.5.3. Intramacronucleata………………………………………………184 7.5.4. Colpodea…………………………………………………………186

7.6. References…………………………………………………………………188

BIBLIOGRAPHY………………………………………………………………………195

xiii LIST OF TABLES

Figure Page

1.1 Taxon sampling within the Colpodea……………………………………………31

1.2 Pairwise distance between collections for species sampled more than once…….32

2.1 Taxon sampling and GenBank numbers used in this study……………………...71

2.2 Morphometric data for Aristertostoma marinum population Framvaren Fjord….72

2.3 Comparative morphology of described representatives of four genera in The Cyrtolophosididae…………………………………………………………...73

3.1 Taxon sampling for mtSSU…………………..………………………………...108

4.1 Isolates newly collected for this study………………………………………….130

4.2 Newly sampled loci……………………………………………………………..131

xiv LIST OF FIGURES

Figure Page

1.1 Evolution among some morphological characters within the class Colpodea…...33

1.2 SSU rDNA genealogy of the class Colpodea and potential sister classes……….35

2.1 Light microscopy of living (a, b) and protargol impregnated (c) Aristerostoma marinum cells....………………………………………………………………….74

2.2 Scanning and transmission electron microscopy of Aristerostoma marinum……76

2.3 Schematic drawing of Aristerostoma marinum (right lateral view) that combines diagnostic features as revealed by live cell imaging, protargol staining and scanning electron microscopy.…………………………………………………...78

2.4 Ecophysiological tolerance limits of Aristerostoma marinum population Framvaren Fjord………………….………………………….…………………...80

2.5 Schematic drawing of the idealized oral structures (paroral membranes and adoral organelles) of the Cyrtolophosididae……………………………………..82

2.6 Most likely Bayesian SSU-rDNA genealogy of the class Colpodea…………….84

3.1 Mitochondrial SSU-rDNA genealogy of the Colpodea………………………...109

3.2 Concatenated nuclear and mitochondrial SSU-rDNA genealogy of the Colpodea………………………………………………………………………..112

4.1 Concatenated SSU-rDNA/ITS genealogy of the halteriids……………….....…132

6.1 Phylogeny and distribution of sex within the Colpodea…….………………….171

Supplementary Figures

1.1 SSU-rDNA genealogy of the class Colpodea and potential outgroups with support from all nodes indicated…………………………………………………37

3.1 Nuclear SSU-rDNA genealogy of the Colpodea…………..…………………...113

4.1 SSU-rDNA genealogy of the halteriids………………………………………...134

4.2 ITS genealogy of the halteriids…………………………………………………136

xv Appendix

1.A GenBank accessions used in analyses for both previously sequences Colpodea taxa and outgroups…………………………………………………….39

xvi

CHAPTER 1

MOLECULAR PHYLOGENETIC ANALYSIS OF CLASS COLPODEA

(PHYLUM CILIOPHORA) USING BROAD TAXON SAMPLING

Micah Dunthorn,a Wilhelm Foissnerb and Laura A. Katza,c

aGraduate Program in Organismic and Evolutionary Biology, University of

Massachusetts Amherst, USA bFB Organismische Biologie, Universität Salzburg, Austria cDepartment of Biological Sciences, Smith College, USA

1.1. Abstract

The ciliate class Colpodea provides a powerful case in which a molecular genealogy can be compared to a detailed morphological taxonomy of a microbial group. Previous analyses of the class using the small-subunit rDNA are based on sparse taxon sampling, and are therefore of limited use in comparisons with morphologically-based classifications. Taxon sampling is increased here to include all orders within the class, and more species within previously sampled orders and in the genus Colpoda. Results indicate that the Colpodea may be paraphyletic, although there is no support for deep nodes. The orders Bursariomorphida, Grossglockneriida, and Sorogenida are monophyletic. The orders Bryometopida, Colpodida and Cyrtolophosidida, and the genus

Colpoda, are not monophyletic. Although congruent in many aspects, the conflict between some nodes on this single genealogy and morphology-based taxonomy suggests the need for additional markers as well as a reassessment of the Colpodea taxonomy.

1.2. Introduction

Assessment of phylogeny based on morphological characters is limited in many microbial eukaryotes. In most amoebae and flagellates morphology provides little guidance and taxonomic resolution is not rich below the class and ordinal levels. In contrast, phylum Ciliophora Doflein, 1901 is relatively morphologically rich and has a well-described taxonomy (Lynn, 2003; Lynn and Small, 2002). Class Colpodea Small and Lynn, 1981 provides a particularly good opportunity to compare the power of morphology and molecular analyses in reconstructing the phylogeny of ciliates. The

Colpodea is monographed and contains a number of somatic and oral characteristics that were used to establish an extensive classification (Foisser, 1993a). Because previous molecular investigations of the class are based on sparse taxon sampling (Lasek-

Nesselquist and Katz, 2001; Lynn et al., 1999; Stechmann et al., 1998), molecular support for the groups established by Foissner (1993a) remains to be evaluated.

The Colpodea is one of eleven ciliate classes (Adl et al., 2005; Lynn, 2003).

Although its position in the subphylum Intramacronucleata is established (Lynn, 2003), well-supported evidence for the sister class of the Colpodea remains elusive. With current taxonomic sampling, neither morphology nor molecules give convincing or consistent arguments because of homoplasy, low bootstrap support, and problems from both paralogy and rate heterogeneity in protein-coding genes (Lasek-Nesselquist and Katz,

2001; Lynn et al., 1999; Stechmann et al., 1998). The classes Nassophorea,

Oligohymenophorea, Plagiopylea, and Prostomatea are the likely sister-group candidates.

Historically, members of the class Colpodea were placed in disparate groups based on oral structure differences (Foissner, 1993a; Lynn et al., 1999). With Lynn’s

(1976; 1981) structural conservatism hypothesis, somatic kinety (kinetosomes and associated fibers) differences were found to be more a appropriate guide to the deep divisions within the ciliates. The Colpodea were united because of their left kinetodesmal fiber (LKm fiber) (Foissner, 1993a). This fiber extends posteriorly and to the left of the posterior kinetosome of their somatic dikinetids. In contrast, Bardele (1981; 1989) argues against the monophyly of the class because of differences in the presence or absence of particles (ciliary plaques) in the membrane of the somatic cilia: they are present only in one order in the Colpodea (Colpodida) and are absent in the rest of the class.

The Colpodea are a group of primarily terrestrial ciliates (Foissner, 1993a).

Besides the unique LKm fiber, the class Colpodea contains distinctive silverline patterns of regular meshes: ‘colpodid’, with large, rectangular meshes; ‘platyophryid’, meshes divided by median silverline between the kineties, or ‘kreyellid’, with minute irregular meshes (Foissner, 1993a). Members of the Colpodea also have somatic stomatogenesis, where parental oral structures are partially or completely reorganized before new oral structures develop during cell division (Foissner, 1993a; Foissner, 1996). In general, a single ‘germline’ micronucleus is close to the single ‘somatic’ macronucleus; in at least some taxa in order Cyrtolophosidida the micronucleus and macronucleus share an outer membrane of the nuclear envelope (Foissner, 1993a). Sex has only been demonstrated in

Bursaria truncatella and is unreported in the rest of the class (Foissner, 1993a; Raikov,

1982).

Foissner (1993) monographed about 170 species and established an extensive higher-level classification for the Colpodea. Subsequently, new genera and species have been described (Foissner, 1993b; Foissner, 1993c; Foissner, 1994; Foissner, 1995;

Foissner, 1999; Foissner, 2003; Foissner et al., 2002; Foissner et al., 2003). Foissner’s

(1993a) split the Colpodea into two subclasses: one with the order Bryometopida based on a ‘kreyellid’ silverline pattern; with the rest of the orders in another subclass, based on

‘colpodid’ and ‘platyophryid’ silverline patterns. These silverline patterns were later argued to be misleading, as SSU rDNA places the Bryometopida next to order

Bursariomorphida (Lynn et al., 1999). In large part there is agreement over Foissner’s

(1993a) orders and families among other classifications (e.g., Puytorac, 1994), except order Grossglockneriida is lumped with order Colpodida in Lynn and Small (1997;

2002).

Based on morphological characters, Foissner (1993a) offers several hypotheses for relationships among these Colpodea orders (Figure 1.1, Table 1.1). First, Colpodida and Grossglockneriida are sister taxa since they share merotelokinetal stomatogenesis

(complete reorganization of parental oral structures), which is probably the derived condition (Figure 1.1A, character 12). In contrast, the other orders have the possibly plesiomorphic state of pleurotelokinetal stomatogenesis (partial reorganization of parental oral structures) (Figure 1.1A, character 1); this hypothesis is supported in previous molecular analyses (Lynn et al., 1999; Lasek-Nesselquist and Katz, 2001). Second,

Bursariomorphida, Colpodida and Grossglockneriidae form a clade because of the possibly apomorphic equally-spaced rows of oral polykinetids (Figure 1.1A, characters

9), as opposed to the possibly plesiomorphic brick-shaped adoral organelles (Figure

1.1A, character 1); this hypothesis is not supported in previous molecular and morphological analyses (Foissner and Kreutz, 1998; Lasek-Nesselquist and Katz, 2001;

Lynn et al., 1999). Third, Bryophryida, Bursariomorphida, Colpodida and

Grossglockneriida form a clade because of the possibly apomorphic deep vestibulum

(depression in the cell with oral structures) (Figure 1.1A, character 7), as opposed to the possibly plesiomorphic flat vestibulum (Figure 1.1A, character 1).

Using limited taxon sampling with small-subunit rDNA (SSU rDNA), monophyly of the class Colpodea is strongly supported by least-squares (LS) and neighbor-joining

(NJ), and weakly supported by maximum parsimony (MP) analyses in Lynn et al. (1999).

In contrast, with just one additional taxon sampled, Lasek-Nesselquist and Katz (2001) find the class to be paraphyletic, with Nassophorea embedded within it—although support is weak from NJ, MP, and maximum likelihood (ML).

Here we increase taxon sampling of the Colpodea using SSU rDNA sequences, including morphospecies from all seven orders, multiple morphospecies within most orders, and multiple morphospecies in the genus Colpoda. Our aim is to assess the following hypotheses: (1) the class Colpodea is monophyletic, (2) orders within the class

Colpodea are monophyletic, and (3) the genus Colpoda is monophyletic. We will also discuss other features uncovered during characterization of SSU rDNA sequences: two distinct copies of SSU rDNA in one taxon, a group I intron in another, and evidence for sex in the Colpodea taxa sampled here. Furthermore, we discuss alternative hypotheses of morphological evolution based on the SSU rDNA topology. Results from these analyses will further development of a predictive, tree-based framework for the taxonomy of the

Colpodea.

1.3. Materials and methods

1.3.1. Taxon sampling and collection

To reconstruct an SSU rDNA genealogy of the Colpodea 27 collections representing 22 species were sampled for this study (Table 1.1). Most species sequenced were collected from soil, i.e., from non-flooded Petri dish cultures as described in

Foissner et al. (2002). Some were from the water and mud occurring in the tanks of bromeliad plants (Foissner et al., 2003). Cells were either collected from the raw culture

(with other species in the dish) or were isolated into clonal culture (with one to few starter cells). With the addition of GenBank accessions from previous studies (Appendix

1.A), the current sampling includes exemplars from all orders, 15 families, 18 genera, and seven morphospecies in the genus Colpoda. Outgroup selection is based partially on previous analyses.

1.3.2. Identification

Species were identified according to the monograph of Foissner (1993), using live observations and various silver impregnation techniques. The new species collected here—Bursaria sp., Platyophrya-like, Platyophrya sp., Rostrophyra sp., and Sagittaria sp.—will be described in separate papers.

1.3.3. DNA extraction, amplification, cloning and sequencing

Between 10 and 10,000 cells were picked with a micropipette, washed, and placed into DNA lysis buffer. Genomic DNA was extracted using phenol/chloroform following standard protocols (Ausubel et al., 1993) or with a DNeasy Tissue kit (Qiagen, CA).

Genomic DNA was amplified using universal 5’ and 3’ prime SSU rDNA primers

(Medlin et al., 1998) with one of two polymerases. For some species Vent polymerase

(New England BioLabs, MA) was used with the following cycling conditions: 4:00 at

950; 32 cycles of 0:30 at 950, 0:30 at 540, and 2:00 at 720; 10:00 extension at 720. For others Phusion polymerase (New England BioLabs, MA) was used with the following cycling conditions: 0:30 at 980; 36 cycles of 0:30 at 980, 0:15 at 680, 1:30 at 720; 10:00 extension at 720. Amplified products were cleaned with a low-melt gel and Ultrafree-Da columns (Millipore, MA), or with microCLEAN (The Gel Company, CA).

To assess within-sample variation, amplified products were cloned with the PCR-

SMART Cloning kit (Lucigen, WI), or Zero Blunt TOPO kit (Invitrogen, CA). Positive clones were identified by PCR screening with AmpliTag Gold polymerase (Applied

Biosystems, CA), and minipreped using Qiaprep Spin Miniprep kit (Qiagen, CA). Clones were sequenced with the Big Dye terminator kit (Applied Biosystems, CA), using 5’ and

3’ primers as well as two internal primers (Snoeyenbos-West et al., 2002). All sequences were run on an ABI 3100 automated sequencer.

Three samples required further methods. For the Bryometopus pseudochilodon indel found in this study, a 5’ primer (AAA CAG TTA TAG GCA GGC AAT TG) was designed that spanned both sides of the deletion to make sure the sequences containing the deletion were not an amplification artifact. Genomic DNA was amplified with this primer along with the universal 3’ primer, following the above protocol. For Colpoda aspera and Cyrtolophosis mucicola (from Austria), algal contaminant SSU rDNA sequences were removed by enzymatic digestion. Amplified products were cleaned with microCLEAN. Re-suspended DNA was incubated at 370 for three hours with BamH1

(New England BioLabs, MA). The reaction was stopped with microCLEAN, and the

DNA was cloned with the Zero Blunt TOPO kit and sequenced following the above protocol.

1.3.4. Genealogical analyses

Phylotypes were constructed from the consensus of the multiple sequence reads of the cloned products and edited in SeqMan (DNAStar). Pairwise distances for within samples were calculated as uncorrected distances in PAUP* v4.0b8 (Swofford, 2002).

Phylotypes were aligned using Hmmer v2.1.4 (Eddy, 2001), with default settings. The training alignment for model building was all available ciliate SSU rDNA sequences downloaded from the European Ribosomal Database (Wuyts et al., 2004) and aligned according to their secondary structure. The alignment was further edited by eye in

MacClade v4.05 (Maddison and Maddison, 2005), with ambiguously aligned regions and base-pair positions with more than five taxa having a gap masked. Remaining gaps were treated as missing data.

The GTR+I+G evolutionary model was estimated using hLTR in MrModeltest v2

(Nylander, 2004). Maximum parsimony (MP) analyses were carried out in PAUP* v4.0b8 (Swofford, 2002), with all characters equally weighted and unordered. The TBR heuristic search option was used, running ten random additions with MulTree option on.

Maximum likelihood (ML) analyses were carried out in RAxML v2.2.0 (Stamatakis,

2006) running 100 replicates. Support for MP and ML analyses came from 1000 bootstrap replicates using heuristic searches. Bayesian analyses was carried out using

MrBayes v3.2.1 (Huelsenbeck and Ronquist, 2003) with support coming from posterior

probability using four chains and running 10 million generations. Trees were sampled every 100 generations. The first 25% of sampled trees were considered ‘burnin’ trees and were discarded prior to tree reconstruction. A 50% majority rule consensus of the remaining trees was used to calculate posterior probability. Trees were imaged with

TreeView v1.6.6 (Page, 1996).

1.4. Results

1.4.1. Pairwise distances within collections

SSU rDNA sequences from twenty-seven collections representing 22 morphospecies show, for the most part, less than 0.50% average pairwise difference among clones within samples (Table 1.1). Sequences are deposited in GenBank, numbers

EU039884-EU039908. Six clones from Bardeliella pulchra show more variation with an average pairwise difference of 0.94%. Clones from the Bursaria sp. 2 collection contain two different phylotypes that are 0.94% different. The phylotypes of the Ilsiella palustris collection from Brazil are 0.57% different, while in the Hawaiian collection phylotypes are 0.54% different. The levels of within-collection variation are assumed to be a combination of intraspecific variation and experimental error. Contaminant SSU rDNA sequences were found in a few cases; for example: algae in B. pulchra, Bresslauides discoideus, Colpoda aspera, Cyrtolophosis mucicola from Austria, and I. palustris from

Hawaii; fungi in Bryometopus pseudochilodon, and Mykophagophrys terricola; and a tetrahymenid ciliate from Hausmanniella discoidea.

Five species were collected more than once, allowing for some within species comparison (Table 1.2). There is no variation between the two B. discoideus collected from Dominican Republic. Between the Malaysian and Niger Colpoda cucullus collections the average pairwise difference among collections was 0.47%, with no phylotype shared between the sites. Although there is no difference between the two

Brazilian collections of C. mucicola, the Brazilian and the Austrian collections are 1.71% different with no phylotype shared between the countries. The I. palustris collections are

0.64% different and likewise do not share phylotypes between the sites.

1.4.2. Deletion within one SSU rDNA copy in Bryometopus pseudochilodon

Two distinct SSU rDNA sequences were characterized from the B. pseudochilodon collection (Table 1.1). One sequence corresponds to the other full-length

Colpodea sequences found here and from GenBank accessions. The second sequence is almost identical to the first except there is a 642 bp deletion towards the 5’ prime end of the SSU rDNA sequence and there are two nucleotide differences on the 5’ end. The deletion starts at nucleotide position 129 in E. coli (GB# J01695) (Cannone et al., 2002).

There is no evidence of elevated substitutions in the sequence with the deletion. This shorter sequence was uncovered in two separate amplifications using universal SSU rDNA primers, as well as from amplifications using a 5’ primer (see methods) that was designed to span either side of the deletion (data not shown). The deletion spanned multiple regions of the SSU rDNA molecule that are conserved in all extant organisms

(Mears et al., 2002). We hypothesize that the deletion sequence is a macronuclear variant,

which occurred in the process of macronuclear development and has been perpetuated during asexual divisions.

1.4.3. Intron in Cyrtolophosis mucicola

A 427 bp intron was found in all SSU rDNA clones from C. mucicola collected from Austria but not the C. mucicola collected from Brazil. The start residue is T and the ending residue is G, which is consistent with group I introns. Blast results also point to this sequence being a group I intron (E value= 1e-16 with the group I intron in Fulgio septica, GB# AJ555452.1; E value= 5e-13 with the group I intron in Acanthamoeba sp.,

GB# EF140633.1). There is no evidence for a homing endoculease gene in the intron.

The insertion position of this intron in the SSU rDNA molecule corresponds to nucleotide

516 in E. coli (GB# J01695) (Cannone et al., 2002), which is a hotspot for group I intron insertions (Jackson et al., 2002).

1.4.4. SSU rDNA genealogy of the Colpodea

After a preliminary analysis using multiple exemplars from all eleven ciliate classes, only Colpodea sequences and close outgroups were chosen for more detailed analyses. The potential sister classes in this analysis as determined in the preliminary global ciliate analysis are the same as in previous studies: Nassophorea, Plagiopylea,

Prostomatea, and Oligohymenophorea (data not shown). One phylotype from each sampled species was used in the alignment, except two representatives of C. mucicola

(because they may underlie two species, see below) and Bursaria sp. 2 (because the other

Bursaria sequences are relatively close).

The final SSU rDNA alignment used for comparing the morphological hypotheses of the Colpodea and its subgroups includes 59 sequences and has a length of

1582 unmasked nucleotides, of which 219 are parsimoniously informative. The most parsimonious tree from the MP analysis is 3349 in length, with a Consistency index of

0.3842, and a Homoplasy index of 0.6157. The most likely tree from the ML analysis has a log likelihood of -17450.098, while the most likely tree from the Bayesian analysis has a log likelihood of -17445.169.

Here we present only the most likely Bayesian tree with node support from all three methods (Figure 1.2, see Supplementary Figure 1.1 for all node support values).

The topologies of the MP- and ML-derived genealogies are mostly congruent with the

Bayesian topology, except in three places. First, in the MP and ML analyses

Cyrtolophodidia II (see below) is basal to the rest of the

Colpodea+Oligohymenophorea+Plagiopylea+Prostomatea with no bootstrap support, and a paraphyletic Nassophorea is basal to this group with no bootstrap. Second, in the MP and ML analyses Bryometopus pseudochilodon is basal to the rest of its order with no bootstrap support, while in the Bayesian tree B. sphagni is basal. Third, in the MP and

ML analyses the order Grossglockneriida forms an unsupported clade with Colpoda aspera, C. steinii, Chain-forming colpodid, and Hausmaniella discoidea, while in the

Bayesian tree it does not.

In our analyses, there is no support for the monophyly of the Colpodea: monophyly of the class is weakly rejected by all three methods based on tree topologies and support values. Furgasonia and Obertrumia (both in the class Nassophorea) fall out sister to part of the order Cyrtolophosidida with no support from all three methods (- MP

bootstrap/- ML bootstrap/- Bayesian posterior probability; support >50% or 0.5 is shown as ‘-’). Changing the number of outgroup classes does not significantly alter this nonmonophyletic topology as the deep nodes are not well resolved anyways and there is no support for any class to be sister to the Colpodea. The rest of the Colpodea forms a monophyletic clade with weak support from MP and ML but with high support from

Bayesian analysis (53/56/0.99).

Support for relationships among the outgroups varied by method. The class

Prostomatea is paraphyletic with only moderate support from Bayesian analysis (-/-

/0.90), with the genus Coleps sister to a well-supported monophyletic class Plagiopylea

(100/100/1.00). The monophyly of Oligohymenophorea is moderately to highly supported in ML and Bayesian analyses (59/71/1.00). The clade containing Prostomatea,

Plagiopylea and Oligohymenophorea is moderately to highly supported by ML and

Bayesian analysis (53/-/1.00).

Monophyly of the morphologically defined groups could be assessed with our single gene tree for every order within the Colpodea, except Bryophryida as only one morphospecies was sampled for this order. Sorogenida, with two genera, is monophyletic with full support (100/100/1.00). Bursariomorphida, with four taxa, is monophyletic also with high support (93/89/1.00). Grossglockneriida, with two genera, is likewise monophyletic with moderate to high support (90/82/1.00).

Order Cyrtolophosidida, with six sampled genera, is not monophyletic. The genus

Cyrtolophosis falls sister to the order Colpodida with moderate to high support

(86/96/1.00). The remaining Cyrtolophosidida genera (Ottowphrya, Platyophrya-like,

Platyophrya, Rostrophrya, and Sagittaria) form a paraphyletic group, with order

Sorogenida nested within it, at the base of the class with high support (100/99/1.00).

Order Bryometopida, with one genus and three species sampled, is not monophyletic with full support from all three methods (100/100/1.00). Order

Bursariomorphida is nested within this order, and it is sister to Bryometopus pseudochilodon with full support (100/100/1.00). To determine whether this topology is spurious due to the GenBank accession for Bryometopus sphagni missing about 500 bp from the 5’ end, the Bryometopida and Bursariomorphida sequences were realigned (with

C. magna and C. mucicola as outgroups) minus the 5’ end; the same topology was found with this alignment (data not shown). Order Colpodida is not monophyletic with high support (97/93/1.00), containing orders Grossglockneriida and Bryophryida. B. pulchra is sister to Notoxoma parabryophryides (order Bryophryida) with moderate to high support

(87/94/0.98).

Monophyly of the genus Colpoda was assessed in this molecular analysis using eight morphospecies within the genus and numerous close outgroups. Colpoda is not monophyletic in the SSU rDNA genealogy with moderate to full support (73/72/1.00).

Most Colpoda species form a sister group to the Grossglockneriida with no support from any method (-/-/-). B. vorax, B. discoideus, and Colpoda henneguyi form a clade with moderate to high support (89/84/1.00). This clade is in turn sister to most of the remaining Colpoda species with moderate to high support (72/80/0.97). To determine if the topology of the Colpoda phylotypes is robust, only Colpodida, Grossglockneriida, and Bryophryida phylotypes were realigned and remasked for a separate analysis; overall, the resulting ingroup topology is concordant with the full class analysis (data not shown).

1.5. Discussion

1.5.1. Comparisons between morphology and molecules

Here we compare the morphologically-based classification and well-supported

SSU rDNA nodes. Furthermore, we evaluate the possible evolution of morphological characters in light of the SSU rDNA genealogy.

1.5.1.1. The class: We find no molecular support for the monophyly of the class

Colpodea based on analyses of SSU rDNA sequences (Figure 1.2). Conversely, the nonmonophyly of the class (with part of the class Nassophorea being sister to part of the order Cyrtolophosidida) is not well supported either. Similarly, the Nassophorea is also not monophyletic with respect to the Colpodea, though with no support. The nonmonophyletic relationships of the Colpodea with respect to the Nassophorea should not be given much weight, as there is neither support for this relationship nor for the

Nassophorea even being sister to the Colpodea. The SSU rDNA genealogy here provides little support for class-level relationships within the subphylum Intramacronucleata in general, as seen elsewhere (Lynn, 2003).

These results do not pose a serious challenge to Lynn’s (1976; 1981) structural conservatism hypothesis given the limited support at deep nodes. On the other hand, these results do challenge Bardele’s (1981; 1989) use of ciliary plaques in his argument that the members in the Colpodea are not closely related.

1.5.1.2. The orders: Molecular support for monophyly could be assessed for all orders within the Colpodea except the Bryophryida. The SSU rDNA genealogy presented here

does support much of the morphologically-based classification of the Colpodea, although there is some discordance at the ordinal level between morphology and molecules

(Figure 1.2).

The order Cyrtolophosidida is polyphyletic. Cyrtolophosidida I, containing the genus Cyrtolophosis, falls away from Cyrtolophosidida II, containing the most recent common ancestor of Sagittaria and Platyophrya and all of its descendants plus the order

Sorogenida. This nonmonophyly of the Cyrtolophosidida suggests the need for a reevaluation of the character that was used to establish this group. Cyrtolophosidida was circumscribed based on the shared outer membrane of the nuclear envelope of the micronucleus and macronucleus (Foissner, 1985; Foissner, 1993a). This character, however, has only been confirmed with transmission electron microscopy for six species:

Aristerostoma marinum (Detcheva and Puytorac, 1979), Cyrtolophosis mucicola

(Detcheva, 1976; Didier et al., 1980), Platyophrya sphagni (Kawakami, 1991),

Platyophrya spumacola (Dragesco et al., 1977), Pseudocyrtolophosis alpestris (Foissner,

1993a), and Woodruffides metabolicus (Golder, 1976). Njine (1979) states that nuclei in

Kuklikophrya ougandae share an outer membrane (and presents a drawing of a stained cell showing this), but does not present an electron micrograph. Platyophryides latus is drawn with a shared outer membrane by Dragesco and Dragesco-Kernéis (1979), but

Puytorac et al. (1992) show that the membranes are separate with their transmission electron micrographs. Foissner (1993a) argues that two taxa, Sagittaria australis and

Woodruffia australis, have the shared outer membrane because of their thick silver- stained membranes. On the other hand, Díaz et al. (2000) show separate outer nuclear membranes in Cyrtolophosis elongata. Hence, the shared outer membrane of the nuclear

envelope of the micronucleus and macronucleus is not only a weak character for the

Cyrtolophosidida, but also one whose distribution is neither well known nor confirmed

(Figure 1.1B, character 4). Future transmission electron microscopy studies are much needed to confirm the presence or absence of this character in other species. Foissner et al. (2002) suggest those species with a separate outer micronucleus and macronucleus membrane can be transferred to the clade Plesiocaryon or into the order Sorogenida (as was done with Ottowphrya).

There are morphological differences between the two Cyrtolophosidida groups. In

Cyrtolophosidida I, there are two segments in the paroral (right oral) membranes, the anterior bearing tuft-like cilia (the unique feature of its family) (Figure 1.1B, character

15). Only one paroral segment is present in taxa in Cyrtolophosidida II (Figure 1.1B, character 14). These groups also differ in the presence of non-ciliated kinety on the right margin of the adoral organelles in Cyrtolophosidida I (and its family), which is absent in

Cyrtolophosidida II (Figure 1.1B, character 16).

Although originally placed with the haptorid ciliates (Bradbury and Olive, 1980), the close relationship between the Sorogenida and the Cyrtolophosidida was soon recognized morphologically (Bardele et al., 1991; Foissner, 1985; Small and Lynn,

1981). This relationship was confirmed in a previous SSU rDNA analysis (Lasek-

Nesselquist and Katz, 2001) and the SSU rDNA topology presented here. The Sorogenida was originally separated from the Cyrtolophosidida because it lacked the shared outer membrane of the nuclear envelope of the micronucleus and macronucleus (Foissner,

1985; Foissner, 1993a)—although this character maybe is weak (see above)—and because of its slime mold-like aerial sorocarp in one life history stage (Fig. 1.1, character

5). Like the Cyrtolophosidida, the Sorogenida has brick-shaped organelles on the left slope of the vestibulum and pleurotelokinetal stomatogenesis (Figure 1.1B, character 1)

(Foissner, 1993a). The SSU rDNA genealogy suggests that the aerial sorocarp of

Sorogena may represent a complex apomorphy arising from within Cyrtolophosidida II.

The order Bryometopida is paraphyletic in relation to the monophyletic

Bursariomorphida in the SSU rDNA genealogy. The close relationship between

Bryometopida and Bursariomorphida was also found by Foissner and Kreutz (1998) and

Lynn et al. (1999). Although these two orders differ in their silverline pattern

(Bryometopida having ‘kryellid’ to ‘platyophryid,’ Bursariomorphida having ‘colpodid’), taxa in these two orders share an apical oral opening, a ventral cleft, conspicuous adoral organelles, and an emergence pore in their cysts (Foissner and Kreutz, 1998; Foissner, pers. obs.).

The order Colpodida is paraphyletic in our molecular analyses, though support is limited at many nodes. That the Grossglockneriida was close to the Colpodida has been proposed as they share the unique (in the Colpodea) merotelokinetal stomatogenesis

(Figure 1.1B, character 12), colpodid silverline pattern (Figure 1.1B, character 6), and a simple oral polykinetid (Aescht et al., 1991; Foissner, 1993a). These two orders are even lumped together in some classifications (Lynn and Small, 1997; Lynn and Small, 2002).

The question remained just how they were related: the SSU rDNA genealogy here suggests that the Grossglockneriida falls within the Colpodida, not sister to it. The position of Bryophryida within the Colpodida has not been hypothesized as the

Bryophryida has a platyophryid silverline pattern and brick-shaped organelles on the left

vestibulum (Figure 1.1B, character 1). The Bryophryida and Colpodida do, though, share a deep vestibulum (Foissner, 1993a).

The use of differences in the type of division seems to be helpful at the ordinal level. As suggested by Foissner (1993a): pleurotelekinetal stomatogenesis is probably plesiomorphic within the Colpodea. Only orders Colpodida and the Grossglockneriida have merotelokinetal division (Foissner, 1993a). Stomatogenesis is undescribed in

Bryophryida; assuming that its phylogenetic position found here is confirmed in future studies, then it is predicted that its division type should be merotelokinetal. On the other hand, the power of the silverline pattern for use in the systematics of the Colpodea at the ordinal level is debatable. While Foissner (1993a) uses differences in silverlines to help construct a higher-level classification, Foissner and Kreutz (1998) Lynn et al. (1999) argue that this character is sometimes misleading. The results presented here are in agreement with Lynn et al. (1999) on the limitations of the use of silverline patterns at the ordinal level.

1.5.1.3. The genus Colpoda: In our molecular analyses the large genus Colpoda is paraphyletic not only in relation to genera within its own family, but also to other families in its order (Figure 1.2). Most of the relationships among the Colpoda morphospecies in the SSU rDNA tree are not well supported; there is support for

Bresslaua and Bresslauides nesting within the Colpoda. Bresslaua was originally separated from Colpoda based on a difference in vestibulum size (Kahl, 1931). However,

Claff et al. (1941), Foissner (1985; 1993a), and Lynn (1979) find that Bresslaua’s voracious feeding behavior and its left-projecting vestibular wall (as opposed to right-

projecting in Colpoda) are probably better characters to separate the genus from Colpoda.

The SSU rDNA topology suggests that these characters may represent apomorphies arising from within a Colpoda clade. Bresslauides (and its family Hausmanniellidae) also falls within the Colpoda in the SSU rDNA tree. This genus was circumscribed based on the unique semicircular right oral polykinetid that was longer than the left as opposed to being equal in the Colpodidae (Foissner, 1987; Foissner, 1993a). Because Bresslauides is not falling out with the other member of its family (Hausmanniella) sampled here, the character of a semicircularly curved right oral polykinetid may have evolved more than once.

1.5.2. Open questions with some species designations

The level of diversity among some SSU rDNA sequences from the morphospecies collected here suggests possible problems with some circumscriptions. Colpoda magna and C. minima differ little in the SSU rDNA phylotypes, indicating a need for further genetic studies. These morphospecies species differ in size and kinety number, as well as the number of micronuclei, with one in C. minima and 2-16 in C. magna (Foissner,

1993a). The low genetic distance between these two species and the lack of much morphological differences could point to these being nascent but “biological” species.

Alternatively, C. minima and C. magna may represent morphological variation within a single species where a change in micronuclei and kinety number is correlated with size.

The C. mucicola morphospecies may represent two genetic species: there is a putative group I intron in the Austrian collection that is absent from the Brazilian collections, and there is greater than 1% pairwise distance between the Austrian and

Brazilian collections. In contrast, the diversity in SSU rDNA phylotypes for the genus

Bursaria from this study and GenBank accessions (1.31% average pairwise distance) supports the view that there are more than one species in the genus, although some argue for there being only one species.

We do not find the large sequence diversity in our Colpoda morphospecies as does Nanney et al. (1998). In our analysis we find a 2.79% average pairwise distance among the Colpoda morphospecies sampled here and from GenBank accessions, while

Nanney et al. (1998) find an average “slack” value of 31.5% among their Colpoda. There are at least two reasons for this difference. First, our analyses were based on SSU rDNA, while theirs is based on 190 bp of the hyper-variable D2 region of the large subunit rRNA

(LSU rDNA). Second, our analyses of distance used the uncorrected distance method in

PAUP*, while theirs use string analyses in the program PHYLOGEN. Using our distance method, Nanney et al.’s (1998) data show an average pairwise distance of 20.73% for the

D2 region of the LSU rDNA (data not shown). Despite the difference in levels of variation between the SSU rDNA and the short variable region of the LSU rDNA, the topology found by Nanney et al. (1998) among their five Colpoda morphospecies is congruent with our analyses (data not shown).

1.5.3. Evidence for sex

Conjugation (ciliate sex) is documented in all ciliate classes (Bell, 1988; Dini and

Nyberg, 1993; Miyake, 1996; Sonneborn, 1957). In the Colpodea conjugation is only known in B. truncatella even though over the decades researchers have looked for conjugation in other species but have yet to observe it (Foissner, 1993a; Raikov, 1982).

There are a few reports of possible conjugation in some species of Colpoda; because nuclear division or exchange was not shown these observations are possibly of

“pseudoconjugation,” where exchange of nuclei does not occur (Foissner, 1993a).

Assuming that Colpodea species behave genetically in a way similar to other eukaryotes, we could predict that if the Colpodea were asexual, allelic variation would be high within species (Mark Welch and Meselson, 2000; Normark, et al. 2003). The low allelic values within most collections sampled here suggest that the Colpodea species are indeed having sex albeit covertly. There is an important caveat in this statement in that the number of clones sequenced per morphospecies in this study is relatively low (1-7 clones) and we could have missed some variation. The results here are in opposition to

Bowers et al. (1998), who present isozyme evidence for asexuality for three Colpoda species. Although cryptic sex is consistent with the low allelic values found here, further evidence of conjugation is much needed to confirm sex within the Colpodea beyond B. truncatella.

1.5.4. Group I intron in Cyrtolophosis mucicola

While group I introns are widespread in microbial eukaryotes (Bhattacharya et al.,

1996; Haugen et al., 2003; Haugen et al., 2005; Snoeyenbos-West et al., 2004; Wikmark et al., 2007), the putative group I intron found in the Austrian C. mucicola morphospecies is the fourth identification of this type of intron in ciliates. The other known species with group I introns are: Tetrahymena thermophila (Grabowski et al., 1981), Acineta sp.

(Snoeyenbos-West et al., 2004), and Tokophrya lemnarum (Snoeyenbos-West et al.,

2004). Undoubtedly there remain more of these introns to be uncovered in future

sequencing projects of the various ciliate groups. We suggest that the intron in the

Austrian Cyrtolophosis mucicola is a product of a recent horizontal transfer into the SSU rDNA locus, as group I introns are known to be mobile over relatively short evolutionary time scales (Haugen et al., 2005; Simon et al., 2005) and because it was not found in other isolates of the species or other Colpodea sequences.

1.5.5. Reconciling morphology and molecules in the Colpodea

In large part morphology and the SSU rDNA genealogy agree in the hypothesized relationships within the ciliate class Colpodea, although the paraphyletic relationships among previously hypothesized closely related taxa was unexpected (Figure 1.1). The

SSU rDNA genealogy is based on a single gene and may not follow the actual species phylogeny (e.g., Doyle, 1992; Maddison, 1997). Further tests using other loci are needed to confirm the areas where there is discordance between morphology and molecules.

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Table 1.1. Taxon sampling within the Colpodea. Species were identified using silver impregnation by W. Foissner. Type and voucher material of the new species and the newly investigated populations are deposited at the Oberoesterreichische

Landesmuseum in Linz (LI), Austria. nc - non-clonal culture, c- clonal culture, npc - non-pure culture, pc - pure culture.

Table 1.2. Pairwise distance between collections for species sampled more than once. Pairwise Taxon Collection site distance (%) Bresslauides discoideus Dominican Republic 1 and 2 0 Colpoda cucullus Malaysia and Niger 0.47 Cyrtolophosis mucicola Brazil 1 and 2 0 Cyrtolophosis mucicola Brazil 1 and Austria 1.71 Ilsiella palustris Brazil and Hawaii 0.64

Figure 1.1. Evolution among some morphological characters within the class Colpodea.

(A) Hypotheses of relationships among orders and morphological character evolution modified from Foissner (1993a), where some characters are removed. (B) Possible alternative evolution of characters mapped out on the SSU rDNA gene tree found here; the deeper nodes in the Colpodea are not well supported and are thus shown as a polytomy. The character are: 1) Lkm fiber, pleurotelokinetal stomatogenesis, brick- shaped adoral organelles, flat vesitibulum, and ‘kreyellid,’ ‘platyophryid,’ or ‘colpodid’ silverline pattern; 2) ‘kreyellid’ silverline pattern; 3) ‘platyophrid’ or ‘colpodid’ silverline pattern; 4) shared micronuclear and macronuclear outer membrane of the nuclear envelope; 5) aerial sorocarps; 6) ‘colpodid’ silverline pattern; 7) deep vestibulum, 8) paroral formation with radial ciliary fields; 9) equidistantly spaced adoral organelles; 10) conjugation; 11) emergence pore in resting cysts; 12) merotelokinetal stomatogenesis;

13) feeding tube; 14) one paroral membrane segment; 15) two paroral membrane segments; 16) postoral pseudomembrane. See text for explanations of characters.

Figure 1.2. SSU rDNA genealogy of the class Colpodea and potential sister classes. The most likely Bayesian tree is shown. Bayesian posterior probability support is shown by differences in thickness of branches. Numerical values from bootstrap support is shown next to the branches as: MP bootstrap/ML bootstrap. Values <50% are shown as ‘-’.

Monophyletic classes and orders are labeled with a solid line, while nonmonophyletic ones labeled with a dashed line. All support values for all nodes are given in

Supplementary Figure 1.

Supplementary Figure 1.1: SSU rDNA genealogy of the class Colpodea and potential outgroups with support for all nodes indicated. The most likely Bayesian tree is shown.

Node support is as follows: MP bootstrap/ML bootstrap/Bayesian posterior probability.

Values <50% are shown as ‘-’. Monophyletic classes and orders are shown with a solid line, while nonmonophyletic ones shown with a dashed line.

Appendix 1.A: GenBank accessions used in analyses for both previously sequenced Colpodea taxa and outgroups. Colpodea: GB# Glaucoma chattoni X56533 Bresslaua vorax AF060453 Glauconema trihymene AY169274 Bryometopus sphagni AF060455 Gruberia sp. L31517 Bursaria truncatella U82204 Haleria grandinella AY007443 Chain-forming colpodid AY398684 Heliophrya erhardi AY007445 Colpoda inflata M97908 Isotricha intestinalis U57770 Colpoda steinii DQ388599 Loxodes magnus L31519 Platyophrya vorax AF060454 Loxophyllum utriculariae L26448 Pseudoplatyophrya nana AF060452 Metopus contortus Z29516 Sorogena stoianovitchae AF300285 Metopus palaeformis AY007450 Nyctotherus ovalis AY007454 Obertrumia aurea* X65149 Outgroups: GB# Ophrydium versatile AF401526 Anophryoides haemophila U51554 Ophryoglena catenula U17355 Anoplophrya marylandensis AY547546 Orthodonella apohamatus DQ232761 Apofrontonia dohrni AM072621 Oxytricha nova X03948 Blepharisma americanum M97909 Paramecium tetraurelia X03772 Caenomorpha uniserialis U97108 Parduczia orbis AY187924 Cardiostomatella vermiforme AY881632 Pleuronema coronatum AY103188 Chilodonella uncinata AF300281 Prorodon teres X71140 Climacostomum virens X65152 Prorodon viridis U97111 Coleps hirtus U97109 Protocruzia sp. AF194409 Coleps sp. X76646 Pseudomicrothorax dubius X65151 Didinium nasutum U57771 Schizocaryum dogieli AF527756 Diplodinium dentatum U57764 Spirostomum ambiguum L31518 Discophrya collini L26446 Stentor roeseli AF357913 Ephelota sp. AF326357 Strombidium purpureum U97112 Epidinium caudatum U57763 Stylonychia lemnae AF164124 Epistylis chrysemydis AF335514 Tetrahymena thermophila X56165 Eufolliculina uhligi U47620 Tokophrya lemnarum AY332720 Euplotes crassus AY007437 Tracheloraphis sp. L31520 Frontonia lynni DQ190463 Trithigmostoma steini X71134 Furgasonia blochmanni X65150 Uronema elegans AY103190 Geleia simplex AY187927 Vorticella campanula AF335518 *In GenBank as Obertrumia georgiana, which is a junior synonym.

CHAPTER 2

PHYLOGENETIC PLACEMENT OF THE CYRTOLOPHOSIDIDAE STOKES,

1888 (CILIOPHORA; COLPODEA) AND NEOTYPIFICATION OF

ARISTEROSTOMA MARINUM KAHL, 1931

Micah Dunthorna, Marion Eppingerb, M. V. Julian Schwarzb, Michael Schweikertc, Jens

Boenigkd, Laura A. Katza,e, and Thorsten Stoeckb

aGraduate Program in Organismic and Evolutionary Biology, University of

Massachusetts, Amherst, Massachusetts, 01003, USA bDepartment of Ecology, University of Kaiserslautern, Erwin-Schroedinger-Strasse 14, D-

67773, Kaiserslautern, Germany cDepartment of Zoology, Biological Institute, University of Stuttgart, Pfaffenwaldring

57, D-67633 Germany dAustrian Academy of Sciences, Institute for Limnology, Mondseestr. 9, A-5310

Mondsee, Austria eDepartment of Biological Sciences, Smith Collage, College Road, Northampton,

Massachusetts, 01063, USA

2.1. Abstract

The ciliate family Cyrtolophosididae Stokes, 1888 contains species that are poorly known from both the morphological and molecular perspectives. To further our understanding of this family we redescribe one species, Aristerostoma marinum Kahl, 1931. Cells in our population have an average in vivo size of 15 x 8 µm. There are six rows of somatic kineties, as well as six dorsal kinetids belonging to sparsely ciliated somatic kineties. The oral apparatus is comprised of a bipartite paroral membrane and four adoral organelles.

The optimal ecological tolerances match those of the environment in which it was collected for pH and O2, but not for salinity and temperature. To further test the phylogenetic placement of the Cyrtolophosididae with increased taxon sampling, we characterize the small subunit rDNA of three morphospecies: A. marinum, Aristerostoma sp. ATCC® Number 50986™, and Pseudocyrtolophopsis alpestris. Unconstrained and constrained molecular analyses support the non-monophyly of the order

Cyrtolophosidida. The family Cyrtolophosididae falls out separately from the rest of its order. We also place haplotypes from previous environmental studies in a phylogenetic context within the class Colpodea.

2.2. Introduction

Like other taxa in the class Colpodea Lynn and Small, 1981, the taxonomic history of the family Cyrtolophosididae Stokes, 1888 is one of shifting classifications.

Members of the group were originally placed in the Frontoniidae by Kahl (1931), and later in the Tetrahymenidae by Corliss (1961). Foissner (1978) used silverline patterns, and Didier et al. (1980), Lynn (1981), and Puytorac et al. (1979) used transmission electron micrographs of kinetid ultrastructure to link this family with other Colpodea.

The Cyrtolophosididae was then placed in the order Cyrtolophosidida with other genera—such as Platyophrya and Sagittaria—in which micronuclei and macronuclei share an outer membrane of the nuclear envelope (Foissner 1985; Foissner 1993).

The Cyrtolophosididae is currently diagnosed with a number of morphological characters: species have a bipartite paroral membrane, “colpodid” silverline pattern, and a non-ciliated kinety on the right margin of their adoral organelles (Foissner 1993).

Morphological variation among the four described genera in the family—Aristerostoma,

Cyrtolophosis, Plesiocaryon, Pseudocyrtolophosis—is not as distinct; Foissner (1993) even suggests that they might need to be synonymized. Most species are relatively small,

20-35 x 15 µm, with a few living in presumably mucocyst-derived tubes (Foissner 1993).

Some of these, like Aristerostoma marinum (part of the focus of this manuscript), lack modern descriptions and silver impregnations.

The phylogenetic placement of the Cyrtolophosididae has recently been questioned. In a molecular analysis of all orders within the Colpodea using small subunit rDNA (SSU-rDNA) sequences, two Cyrtolophosis mucicola sequences branched separately from the rest of its morphologically-defined order (Dunthorn et al. 2008). This

result challenges the use of the shared outer membranes of the nuclear envelope to unite the order Cyrtolophosidida. The non-monophyletic SSU-rDNA topology of the order, though, requires further evaluation with increased taxon sampling.

Here we redescribe A. marinum and a new name-bearing type is designated. We also move sampling beyond the only one sequenced species, C. mucicola, and further test the monophyly of the order Cyrtolophosidida with three previously uncharacterized morphospecies in two genera using SSU-rDNA phylogenetic analyses. Morphological and molecular hypotheses are compared with constrained analyses, and possible issues leading to differences between the morphological and molecular hypotheses are examined. Furthermore, we place GenBank accessions from previous SSU-rDNA environmental surveys in the context of our increased taxon sampling within the

Cyrtolophosididae.

2.3. Methods

2.3.1. Taxon sampling

Three morphospecies in the family Cyrtolophosididae were isolated for this study and SSU-rDNA was sequenced from them (Table 2.1). A. marinum was collected from surface waters of the Framvaren Fjord in southwest Norway (58°09’N, 06°55’E). Pure cultures for a redescription of this species were established using Schmaltz-Pratt medium

-1 -1 -1 (0.01 g K2HPO4 * 3H2O l , 0.1 g KNO3 l , 1.45 g CaCl2 * 2H2O l , 6.92 g MgSO4 * 7H2O

-1 -1 -1 -1 l , 5.51 g MgCl2 * 6H2O l , 0.67 g KCl l and 28.15 g NaCl l ) with heat-inactivated

Klebsiella minuta as a food source. Aristerostoma sp. ATCC® Number 50986™ was

originally collected from a marine environment in the Great Marsh, Delware, USA, where there was gray mud mixed with roots, sand, and clay. This isolate will be examined morphologically elsewhere. Pseudocyrtolophopsis alpestris was collected from litter of a spruce forest in Lambrechtshausen near Salzburg, Austria, by W. Foissner.

With the addition of the two GenBank accessions for C. mucicola and sequences from environmental studies (Table 1), we now have eight exemplars in the

Cyrtolophosididae. Sequences from the rest of the Cyrtolophosidida and from the other

Colpodea orders are from GenBank (Table 1). Outgroup selection was based on previous studies.

2.3.2. Light and electron microscopy

For light microscopy of living and stained cells, we used a Zeiss Axioplan 2.

Protargol impregnation followed Foissner et al. (1999), with the cells fixed in 1 ml

aqueous saturated HgCl2 with 100 µl Bouin’s fluid (cells vol/fixant vol 1:1, 30 min, RT).

Due to cell sensitivity, a high salt concentration in the medium (causing precipitates during processing), and a mucus shell covering the organisms (mucocysts, see Results) other staining and impregnation methods failed (e.g. silver nitrate impregnation with the

Chatton-Lwoff technique and silver carbonate impregnation with the Fernandes-Galiano technique for saltwater ciliates). All images from living, fixed, and stained cells were taken by a QImager® Microcam (Intas, Göttingen, Germany) and QCapture© software

(http://www.qimaging.com). For further image processing we used Adobe Photoshop©

7.0 and ImageJ 1.32 (http://rsb.info.nih.gov/ij/).

For scanning electron microscopy (SEM), cells were processed following Stoeck et al. (2005) with slight modifications for fixation: cells/fixans, 1/1; fixans of osmium tetroxide (2% in artificial seawater [36‰] and ASW following Stetter et al. (1983) for 60 minutes at room temperature). Stubs with fixed and dehydrated cells were coated with gold (Edwards E306) and observed with a Zeiss DSM940A (Carl Zeiss GmbH, Jena,

Germany).

Preparation of cells for transmission electron microscopy (TEM) followed Stoeck et al. (2005) with slight modifications in the fixation procedure: a culture aliquot was first fixed with glutaraldehyde (2.5% final) for 60 min at 4°C. Cells were pelleted and embedded in a 4% low-melt sea-prep agarose (Roth, Karlsruhe, Germany) (Reize and

Melkonian 1989) with 4% osmium tetroxide in ASW for 60 min in order to concentrate and handle the small target cells. Ultrathin sections were investigated with a Zeiss EM10

(Oberkochen, Germany) and documented on a Kodak 4489 film (Eastman Kodak, NY).

2.3.3. Terminology

Terminology follows Corliss (1979), Foissner (1993), and the International Code of Zoological Nomenclature (ICZN 1999). In designating a new type specimen, we follow Foissner (2002) by allowing the new name bearer to be from a different location than the specimen originally described by Kahl (1931).

2.3.4. Autecology

The ecological tolerances of A. marinum towards four parameters (% O2 in the headspace gas, pH, ‰ salinity, and temperature) were experimentally tested in 2-ml

batch incubations. Cell activity was measured by counting moving and/or dividing cells under a dissection microscope within 24, 48, 72, 96 and 168 h periods after inoculation.

The cultures were gradually adapted to higher/lower pH, salinity, and temperature as outlined by Stoeck et al. (2005).

All incubations were inoculated in six 2-ml parallels (six wells on a 24-wellplate,

Greiner, Germany) in chemically adjusted Schmaltz-Pratt medium at room temperature

(with the exception of the temperature experiment). Salinity was changed by the addition

of 1 M NaCl or Volvic™ water. Changes in pH were adjusted by addition of 1 M NaCO3

or 1 M KH2PO4. The cells were inoculated after pH stabilization (24 h). Heat-deactivated

K. minuta were added as a food source at saturated concentration (108-109 cells/ml).

The preferred oxygen regime was tested by incubation of seven 2-ml parallels in

10-ml injection bottles (Ochs GmbH, Bovenden-Lenglern, Germany). These were stored inside a gas-tight 1000 ml glass chamber containing a defined headspace gas composition

(0, 1, 2 or 21% oxygen in N2, 4.0 calibration gas qualities, AirLiquide, Darmstadt,

Germany). Final O2-concentration in the medium was reached after 24 h (t0). Anoxic conditions were established by flushing the medium and incubation vessel with N2

(AirLiquide, Darmstadt, Germany) and by the addition of anaerocult-plates (Merck AG,

Darmstadt, Germany) as an oxygen scavenger. Suboxic and oxic incubations were flushed twice daily with the appropriate calibrated gas. For each gas concentration, we prepared four replicates, one of which was sacrificed after each of the testing periods (24,

48, 72, 96 and 168 h) to count moving and/or dividing cells. All experiments were incubated at room temperature in the dark.

2.3.5. Amplification and sequencing

To extract genomic DNA, 0.5-ml aliquots of a culture or 5-10 individually picked cells were picked with a micropipette, washed, and processed using the protocol for cultured animal cells of the DNEasy Tissue Kit (Qiagen, Hildesheim, Germany). SSU- rDNA was amplified using the universal eukaryotic primers EukA and EukB (Medlin et al. 1988). For A. marinum and P. alpestris, each amplification contained 10-20 ng of

DNA template, 2.5 U HotStar Taq DNA polymerase (Qiagen) in the manufacturer-

provided reaction buffer, 1.5 mM MgCl2, 200 µM of each dNTP, and 0.5 µM of each oligonucleotide primer. The final volume was adjusted to 50 µl with sterile distilled water. The PCR protocol for SSU-rDNA gene amplification consisted of an initial hot start incubation of 15 min at 95 ˚C followed by 30 identical amplification cycles (i.e., denaturing at 95 ˚C for 45 s, annealing at 55 ˚C for 1 min, and extension at 72 ˚C for 2.5 min), and a final extension at 72 ˚C for 7 min. Negative control reactions included

Escherichia coli DNA as a template. The resulting PCR products were cleaned with the

PCR MinElute Kit (Qiagen) and cloned into a vector using the TA-Cloning kit

(Invitrogen, Carlsbad, CA). Plasmids were isolated with Qiaprep Spin Miniprep Kit

(Qiagen) from overnight cultures and PCR-reamplified using M13F and M13R primers to screen for inserts of the expected size (ca. 1.8 kb in case of the SSu-rDNA fragment). For

Aristeristoma sp., Phusion polymerase (New England BioLabs, MA) was used for amplification and the products were cloned following Dunthorn et al. (2008). All clones were sequenced bidirectionally (M13 sequence primers) with the Big Dye terminator kit

(Applied Biosystems, Foster City, CA) on either an ABI 3100 or 3730 automated sequencer.

2.3.6. Genealogical analyses

We determined and edited haplotypes from overlapping sequence reads in

SeqMan (DNAStar, Inc., Madison, WI) or CodonCode Aligner v1.2.4 (CodonCode

Corporation, Dedham, MA). Pairwise distances for within and among samples were calculated as uncorrected “p” distances in PAUP* v4.0b8 (Swofford, 2002). Haplotypes generated here and the environmental sequences from GenBank were placed into the alignment used in Dunthorn et al. (2008), but with most non-Colpodea outgroups removed. The GTR+I+G evolutionary model was selected using hLTR in MrModeltest v2 (Nylander 2004).

Maximum parsimony (MP) and maximum likelihood (ML) analyses were carried out in PAUP* v4.0b8 (Swofford 2002), with all characters equally weighted and unordered. The TBR heuristic option was used to search trees, running ten random additions with MulTree option on. Support for MP and ML analyses came from 100 bootstrap replicates using heuristic searches. ML bootstraps were run on the Beowulf cluster at the University of Missouri St. Louis. Bayesian analyses was carried out using

MrBayes v3.2.1 (Huelsenbeck and Ronquist 2003) with support coming from posterior probability using four chains and running 10 million generations. Trees were sampled every 1000 generations. The first 25% of sampled trees were considered ‘burnin’ trees and were discarded prior to tree reconstruction. A 50% majority rule consensus of the remaining trees was used to calculate posterior probability.

2.3.7. Constrained analysis

In addition to the genealogical analysis above, a ML analysis was carried out with all exemplars in the Cyrtolophosidida constrained to be monophyletic in PAUP* v4.0b8

(Swofford 2002); the particular relationship within the Cyrtolophosidida, though, were not specified. The resulting tree was compared to the unconstrained ML tree using a one- tailed KA test (Kishino and Hasegawa 1989) as implemented in PAUP* v4.0b8

(Swofford 2002).

2.3.8. Rate class analyse

Using the full unconstrained alignment, nucleotide positions were partitioned into eight rate classes using HYPY v0.9b (Kosakovsky Pond et al. 2005). The fastest rate class was removed from the alignment, and explored using Bayesian analysis as above, except running 3,000,000 generations. The second fastest rate class was then also removed and examined likewise.

2.4. Results

2.4.1. Description of the neotype of Aristerostoma marinum Kahl, 1931

While free-swimming, the fast cells spirals, rotating around their longitudinal axis. Cells have no tendency to clump together either while swimming or in the resting state. After a few minutes under the microscope numerous cells attached either to the water surface or to the cover slip with their posterior end. In vivo, the ciliate is 9-23 µm in length (mean 15 µm, n = 29; measurements rounded to the nearest digit) and 4-11 µm

wide (mean 8 µm, n = 29) (Table 2.2). The neotype population of A. marinum has an oval shape tapering towards the anterior end. An oral structure is visible in the anterior third of the cell (Figs. 2b, 2.3), while the posterior end displays a pulsating vacuole (Fig.

2.1A). While swimming it becomes visible that the left dorsal side is flattened. The cell surface displays prominent longitudinal ribs, and on the right lateral side the cortex carries easily recognizable rows of cilia. Only a few irregularly distributed cilia are recognizable on the dorsal side. In vivo the somatic cilia are ca. 4 µm in length. We did not observe any resting stages in any of our cultures. Reproduction occurs by symmetrogenic binary fission (=perkinetal, Fig. 2.1B).

All protargol impregnations are suboptimal (Fig. 2.1C) and only the examination of numerous cells enabled a schematic drawing of protargol impregnated structures (Fig.

2.3). Cells are 10-20 µm in length (mean 15 µm, n = 47) and 6-10 µm wide (mean = 8

µm, n = 47). The oval-shaped macronucleus is located submedian (in the posterior half of the cell) and has a mean diameter of 3 µm. In protargol preparations, the micronucleus is sometimes delimited from the macronucleus as a lighter-colored structure with a mean diameter of 1 µm (Fig. 2.1C). Protargol impregnation does not reveal whether the micronucleus lies within the nuclear membrane. The distribution of extrusomes

(mucocysts) in the cell’s cortex becomes visible (Fig. 2.1C). The oral structure appears subapical, but details cannot be resolved using this impregnation. Protargol impregnated kinetids are displayed in the schematic drawing (Fig. 2.3). The cell is characterized by six rows of somatic kineties. Kinety 1 consists of six dikinetids extending from the anterior end of the cell 2/3 towards the posterior along the longitudinal axis (Fig. 2.3); it is located right laterally. Also the right lateral kineties 2 and 3 consist of eight dikinetids each that

extend along the complete longitudinal axis from anterior to posterior poles (Fig. 2.3).

Kinety 4 (right lateral-dorsal) is composed of four dikinetids and three uniciliated kinetids (which are likely to be dikinetids), running from the anterior end of the cell 2/3 towards the posterior along the longitudinal axis (Fig. 2.3). Kinety 5 (dorsal-left lateral) comprises only two dikinetids located at the anterior end of the cell (Fig. 2.3). Kinety 6

(left lateral - ventral) also consists exclusively of dikinetids (n = 8), which extend along the whole length of the longitudinal axis and abut left lateral the oral apparatus (Fig. 2.3).

Six kinetids are on the dorsal side. They cannot be assigned to any of the six longitudinal kineties but most likely belong to sparsely ciliated somatic kineties (Fig. 2.2A). As we did not succeed in obtaining appropriate transmission electron micrographs of these kineties, we are not able to define if we are dealing with mono- or di-kinetids—further

TEM work is needed.

2.4.2. Electron microscopy

Scanning electron microscopy confirms protargol impregnation results (Fig.

2.2A-C) and reveals details of the subapical oral structure of A. marinum (Fig. 2.2B). A gapless paroral membrane surrounds the triangular oral structure on the right side consisting of 5 anterior dikinetids and 3 adjacent posterior monokinetids (bipartite) (Fig.

2.5). The oral structure’s right margin is bulged and separates the paroral membrane from the oral structure (Figs. 2.2B, 2.3). Four adoral organelles (membranelle 1 to 4) originate from the vestibulum. Membranelle 1 consists of four kinetids and emanates in the upper anterior end of the slightly depressed vestibulum. Three more adoral organelles

(membranelles 2 to 4), each consisting of four kinetids derived from the vestibulum,

comprising two cilia rows each. Two additional dikinetids were located right below the posterior end of the oral structure, which could not be assigned to any of the six somatic kineties (Fig. 2.2A). All dikinetids possess two cilia.

TEM observations (twelve individual cells were analyzed) of our collection of A. marinum show numerous extrusomes (mucocysts) located below the pellicle. These mucocysts are highly sensitive and partly discharged during cell fixation. Mitochondria are characterized by tubular cristae (Fig. 2.2D-2.E). The micronucleus and the macronucleus share an outer membrane of the nuclear envelope; we were not able to clarify the exact organization of this membrane. The nucleolus is located peripherally and clearly visible as a dense round structure.

2.4.3. Autecology

A. marinum is a bacterivore with a preference for smaller bacteria (<1 µm, data not shown). In mixed cultures we did not observe smaller flagellates in the food vacuoles.

Laboratory autecological experiments show that the optimal salinity for cell growth is between 35 and 40‰, with growth ceasing below 17.5 and above 45‰ salinity. Cell growth is highest between pH 7 and 8, with growth ceasing below pH 4 and above pH 10.

At a temperature of 28 °C cell growth is highest, but at temperatures below 12 °C and at

37 °C cell growth stops. A. marinum is an obligate aerobe with highest growth rate when the level of oxygen is 21% in the headspace, with growth ceasing below 1%.

2.4.4. Pairwise SSU rDNA sequence differences within the Cyrtolophosididae

The average pairwise distance among the eight Cyrtolophosididae sequences generated here and from GenBank is 5.413%. The pairwise distance between the two

Aristerostoma spp. collections is 3.99%. The distance between P. alpestris and

HAVOmat-euk43 and LKM63 is 0.65 and 0.889%, respectively.

2.4.5. Genealogical analysis

The SSU-rDNA alignment used for testing the phylogenetic placement of the

Cyrtolophodididae contains 43 sequences, including three new morphospecies sequenced here: Pseudocyrtolophopsis alpestris and two in the genus Aristerostoma. The alignment has a length of 1623 unmasked nucleotides, 347 of which are parsimony-informative. The most parsimonious tree from the MP analysis is 1580 steps in length, with a Consistency index of 0.48, and a Homoplasy index of 0.52. The most likely tree from the ML analysis has a lnL of -10080.85. The most likely tree from the Bayesian analysis has a lnL of -

10109.04 (Fig. 2.6).

With the limited outgroups used for this study, the class Colpodea is monophyletic with weak support from all methods of analysis (57 MP bootstrap/- ML bootstrap/0.88 Bayesian posterior probability; support less than 50% or 0.5 is shown as

‘-’)—but see Dunthorn et al. (2008). Trees generated from each method have largely congruent topologies within the class. In the MP tree, Bryometopus pseudochilodon is basal to its order (plus the order Bursariomorphida), while in the ML and Bayesian trees

Bryometopus sphagni is basal. In the Bayesian tree the chain-forming colpodid+Colpoda steinii and the order Grossglockneriida form a clade, although not supported, while in the

MP and ML analysis these taxa form a polytomy with Colpoda aspera and

Hausmanniela discoidea.

The order Cyrtolophosidida falls out in two groups with moderate to well supported intervening nodes between them. Cyrtolophosidida I contains all exemplars from the family Cyrtolophosididae and the included environmental samples available from GenBank, with strong to full support from all methods (97/100/1.00).

Cyrtolophosidida II contains the remaining exemplars from the order Cyrtolophosidida with the order Sorogenida embedded within them, and receives full support from all methods (100/100/1.00).

Relationships among the sampled Cyrtolophosididae are for the most part well resolved. Both Aristerostoma sequences form a clade with full support from all methods

(100/100/1.00). The genus Aristerostoma is sister the rest of the Cyrtolophosididae with high to full support from all methods (92/87/1.00). Cyrtolophosis mucicola is in turn sister to the Pseudocyrtolophosis nana and the environmental samples with low support from all methods (-/64/0.67).

2.4.6. Comparisons of hypotheses

To compare the morphological hypothesis of Foissner (1993) (where the

Cyrtolophosidida is monophyletic) with that of the SSU-rDNA gene tree estimated here

(where it is not) the likelihood between the alternative hypotheses was examined. The

ML genealogy from the constrained analysis where the Cyrtolophosidida was forced to me monophyletic has a lnL of -10221.46 (tree not shown). This likelihood values is

140.61 less the non-constrained ML genealogy, and is rejected by the KA test (P value <

0.000) in favor of the non-constrained tree. This indicates further support for the non- monophyly of the Cyrtolophosidida

2.4.7. Long-branch attraction

Nucleotides were partitioned into eight rate classes to test the possibility that the non-monophyletic topology of the Cyrtolophosidida is a spurious result due to unequal rates of mutation. If long-branch attraction is effecting the full dataset, then subtracting the fastest evolving nucleotide sites should remove its effect.

The SSU-rDNA alignment with the fastest rate class removed has 1462 unmasked nucleotides, 134 of which are parsimony-informative; the most likely tree from the

Bayesian analysis has a lnL of -5717.45 (data not shown). The SSU-rDNA alignment with the fastest and second fastest rate classes removed has 1256 unmasked nucleotides,

35 of which are parsimony-informative; the most likely tree from the Bayesian analysis has a lnL of -2769.44 (data not shown). In both of these trees much of the structure of the topology is lost among and within clades; however, the Cyrtolophosididae does not branch next to, nor nests within, the other Cyrtolophosidida taxa within either one. These results support the view that the non-monophyletic topology of the Cyrtolophosidida is not the result of long-branch attraction.

2.5. Discussion

2.5.1. Neotypification and emended diagnosis of Aristerostoma marinum Kahl 1931

1931 Aristerostoma marinum Kahl, Tierwelt Dtl., 21:340

1979 Aristerostoma marinum - Detcheva & Puytorac, Annls. Stn. Limnol. De Besse,

13:247

1993 Aristerostoma marinum Kahl 1931 – Foissner, Protozoenfauna, 4/1:557 (revision)

Reference to the Neotype: Individual specimen marked with a circle on slide 1 at the collection of microscopic slides of the Biology Center at the Upper Austrian Museum of Natural History (Linz, Austria), storage code 2007/580-582.

Neotype material: Neotypified from brackish surface waters of the Framvaren

Fjord in southwestern Norway (58°09’ N, 06°55’ E) for the following reasons: (i) no type material is available and no type location has been defined; (ii) the existing descriptions are decisively incomplete; (iii) the genus has a proposed subjective junior synonym

(Foissner 1993); (iv) there are several similar species whose identity is threatened by the species to be neotypified; (v) new preparations (“neotype slides”) are of a quality allowing the specific features to be clearly recognized. The sample from which we isolated A. marinum was taken with a Niskin bottle in a depth of ca. 3 m. At the time of sampling temperature of the surface water was 16 °C, salinity was 15‰ and the water was saturated with oxygen. Nutrient measurements indicated oligotrophic conditions. The species was isolated directly from the natural sample without prior enrichment.

2.5.2. Emended diagnosis

Size in vivo 9 – 23 x 4 – 11 µm, not contractile, oval shape, tapering anteriorly, 1 spherical-elliposid macronucleus located in the centre of the cell, 1 smaller spherical- ellipsoid micronucleus, both nuclei share an outer membrane of the nuclear envelope, extrusomes (mucocysts) hardly recognizable in vivo but impregnate well with protargol,

6 rows of somatic kineties, oral apparatus subapical located anteriorly, oral aperature triangular, four oral membranelles, bipartite paroral membrane on right slope of vestibulum, consists of five anterior dikinetids and three uniciliated kinetids (although they may be dikinetids), 1 contractile vacuole is located posteriorly, no resting stages observed.

2.5.3. Occurrence and Ecology

The type location is not known, but is assumed to be the German Sea coast near

Hamburg (Foissner 1993). Kahl (1931) found A. marinum in infusions (location not defined) and only mentions that it is fairly common and sometimes occurs in high abundances. Detcheva (1982) reported A. marinum from the Bulgarian coast of the Black

Sea where it frequently appeared in high numbers. We here define the Framvaren Fjord in Norway as neotype location. The salinity range of the locations where the Detcheva

(1982) found A. marinum was 1 to 18‰. The salinity of the surface waters in the

Framvaren Fjord, where we found A. marinum, was 15‰. Interestingly, we did not observe growth of our A. marinum population in laboratory experiments below 17‰. At this point we lack an explanation for the discrepancy of the salinity of the populations natural habitat and the laboratory experiments. In contrast, optimal growth at pH 7-8

under laboratory conditions matches the pH of the natural habitat (pH 7.2). Optimum growth temperature under laboratory conditions was between 20 and 31 °C.

Temperatures above 20 °C are only found in the summer month in the surface waters of the Framvaren Fjord. It is interesting that the laboratory cultures did not survive temperatures below 12 °C as the natural habitats temperature may drop below 5 °C in the winter months (data unpublished). As we did not observe that this ciliate is capable to form resting stages; thus, it remains an open question how and where these organisms survive winter temperatures in Norwegian waters.

2.5.4. Comparison with original descriptions and related species

The genus Aristerostoma Kahl, 1926 (order Cyrtolophosidida Foissner, 1978, family Cyrtolophosididae Stokes, 1888) is diagnosed as very small, laterally flattened and completely ciliated (Foissner 1993). As pointed out by Foissner (1993), the currently applied diagnostic characters are based on the incomplete description by Kahl (1931).

The genus is hardly distinguishable by light microscopy from Cyrtolophosis and

Pseudocyrtolophosis, the other two genera within the Cyrtolophosididae. It was suggested that Pseudocyrtolophosis may in fact be a junior synonym of Aristerostoma, or both may in turn be a junior synonym of Cyrtolophosis (Foissner 1993). A major reason for an uncertain species identification is a lack of information about the infraciliature leading to an obscured general morphology of Aristerostoma (Foissner 1993).

One marine and one freshwater species have been described in Aristerostoma by

Kahl (1926; 1931). Both of the taxa lack type material and modern silver stains. While our findings contrast the description of A. minutum (habitat, body shape, longitudinal

rows, see Foissner 1993) they agree with the rudimentary morphological description of A. marinum by Kahl (1931) (Table 2.3) and the ultrastructural description (nuclear and oral structures) of Detcheva and Puytorac (1979) as summarized in Foissner (1993). This is the position and shape of the macronucleus (spherical, in the centre of the cell) and the position of the contractile vacuole (near the posterior end). The oral apparatus is in the anterior third and bordered on right by a paroral membrane. On the opposite side of the oral apparatus four adoral organelles that are composed of two ciliary rows each.

Our analyses revealed the following additional characters of the oral apparatus: (i) the first oral membranelle (membranelle 1) consists only of four monokinetids (Fig. 2.4);

(ii) in contrast to former descriptions (see Foissner 1993) the paroral membrane is not composed of only dikinetids, but of five anterior dikinetids and three posterior monokinetids and thus, is bipartite (Fig. 2.2b, 2.3) (iii) the bipartite paroral membrane does not display a gap. Thus, it is very similar but not identical to other

Cyrtolophosididae genera (Fig. 2.4). For example, the gap between the anterior and posterior segment of the paroral membrane is highly variable in Cyrtolophosis (Fig. 2.4, see Fig. 215b in Foissner 1993) and in Pseudocyrtolophosis about two thirds of the paroral membrane is non-ciliated (Fig. 4, Foissner 1993). In Plesiocaryon elongatum (=

Balantiophorus elongatus Schewiakoff, 1892, = Cyrtolophosis elongata (Schewiakoff,

1892) Kahl, 1931) the bipartite paroral membrane consists of an anterior row of six or seven pairs of kinetosomes whereas the posterior most is a row of five single kinetosomes both being separated by a distinctive gap (see Fig. 4 and Fig. 1b in Diaz et al. 2002).

Furthermore, the posterior portion of Plesiocaryon terricola is composed of an average of four widely spaced, barren monokinetids appearing as minute thickenings in vivo (Fig.

2.4, Foissner et al. 2002). Because of only rudimentary descriptions of A. minutum, at this point we are not able to distinguish A. marinum from A. minutum based exclusively on the oral structure. However, A. marinum is clearly distinct from other genera within the

Cyrtholophosididae regarding the structure of the oral apparatus.

In contrast to Kahl’s population (30 µm), our type material is in average half as long (15 µm, Tables 2 and 3). Decheva and Puytorac (1979)—who did not provide a species epithet—give a length of about 20 µm. A cell width is given in neither of these earlier descriptions. Thus, we add this character to the species diagnosis (9-23 µm in vivo, 10-20 µm after protargol). In his first description, Kahl pointed out that he was not able to see the dorsal infraciliature under light microscopy. However, in a schematic drawing the author shows six longitudinal rows on the right lateral side. Using protargol impregnation and scanning electron microscopy, we can clearly identify six kinety rows with their detailed infraciliature (see Figs. 2.2. 2.3). This serves as an additional criterion for the identification of this species, as all other members of the family Cyrtolophosididae possess 8-10 somatic kinety rows (Table 2.3). The detailed infraciliature of A. minutum is unknown but seems to have more longitudinal kineties (at least eleven according to

Kahl’s (1931) schematic drawing).

We confirm the observation of Detcheva and Puytorac (1979) that the macronucleus and the micronucleus share an outer membrane of the nuclear envelope in the genus Aristerostoma. While they did not identify their isolate down to species, here we show that this character state occurs in A. marinum. Since it is not know how widespread the shared outer membrane is distributed in the Cyrtolophosidida, it is important that each species be investigated using the SEM (Dunthorn et al. 2008). Like

the other species in the genus, we still lack detailed descriptions of division morphogenesis in A. marinum, which will have to be investigated elsewhere.

2.5.5. Neotype specimens

Two neotype slides with protargol impregnated specimens have been deposited at collection of microscopic slides of the Biology Center at the Upper Austrian Museum of

Natural History (Linz, Austria) (storage code 2007/580-582). Live cultures are available from the authors. A circle on slide 1 marks an individual cell that designates the name- bearing type.

2.5.6. Pairwise distance between Aristerostoma morphospecies

In this study we sequenced two different Aristerostoma populations: one from the

ATCC, which is an unidentified species; and one isolated from the Framvaren Fjord in

Norway, which we identified as A. marinum. As the SSU-rDNA sequences of both taxa have a pairwise difference of 4.285% they may represent two different species. Thus, future efforts are in order to characterize the ATCC population in detail. This ATCC population is unlikely to be the other known species in the genus, A. minutum, which is from freshwater.

2.5.7. Phylogenetic placement of the order Cyrtolophosidida

Increased taxon sampling using three new morphospecies sequenced here does not support the monophyly of the order Cyrtolophosidida (Fig. 2.6). Instead, the order divides into two groups that have distinctive oral morphologies. Genera in the clade

Cyrtolophosidida I, containing all Cyrtolophosididae sequences, have a non-ciliated kinety on the right margin of the adoral organelles and the oral apparatus is located sub- apically. Genera in the group Cyrtolophosidida II, which contains the remainder of the order plus the order Sorogenida, lack a non-ciliated kinety and the oral apparatus is located at the apical pole (Dunthorn et al. 2008; Foissner 1993).

To further test the monophyly of the Cyrtolophosidida, we compared Foissner’s

(1993) morphologically-based hypothesis of the Cyrtolophosidida with the topology of the SSU-rDNA maximum likelihood tree found here. An ML analysis of the SSU-rDNA sequences with the non-constrained topology was significantly more likely than the topology constrained to be monophyletic according the KA test. Like the increased taxon sampling discussed above, this comparison also does not support the monophyly of the

Cyrtolophosidida.

2.5.8. Reconciling morphology and molecules

Here, we focus on arguments presented by Foissner et al. (2004) to discuss why there is disagreement between the morphological classification of the Cyrtolophosidida by Foissner (1993) with that of the SSU-rDNA topology found here.

SSU-rDNA sequences are misidentified. The sequences for Cyrtolophosis from

Dunthorn et al. (2008) could be from another ciliate. However, we argue that we can reject this argument as the other sequences from the family Cyrtolophosididae here more closely match the Cyrtolophosis sequences than any other ciliate.

SSU-rDNA sequences are paralogs. Paralogs for protein-coding loci seem to be rampant within ciliates (e.g., Israel et al. 2002; Katz et al. 2004). However, we argue

against this as divergent SSU-rDNA paralogs have yet to be reported within ciliates (or at least ones in which there has not been concerted evolution).

Insufficient taxon sampling. Insufficient taxon sampling can lead to spurious results (Cummings and Meyer 2005; Graybeal 1998; Hedtke et al. 2006; Hillis 1998;

Hillis et al. 2003; Poe and Swofford 1999). We argue that we can reject this argument as we now have eight sequences from the Cyrtolophosididae (representing three of the four genera in the family) and the same non-monophyletic topology is recovered as in

Dunthorn et al. (2008), which only had two sequences from the same morphospecies.

Long-branch attraction. Heterogeneous rates of evolution among branches can lead to spurious results (Felsenstein 1978). All methods of analyses suffer from this problem to one degree or another (Hendy and Penny 1989; Huelsenbeck and Hillis 1993;

Kolaczkowski and Thornton 2004), although the extent of statistical inconsistency in real datasets is questionable (Anderson and Swofford 2004; Bergsten 2005; Siddall and

Whiting 1999). We argue that we can reject long-branch attraction for our SSU-rDNA dataset for two reasons. First, a visual inspection of the genealogy does not show any protruding single or paired long-branches. Second, successively removing the fastest and second fastest nucleotide sites still produced the same non-monophyletic topology.

Gene tree vs. species tree. The topology of any gene genealogy may not accurately reflect the actual species phylogeny (Doyle 1992; Doyle 1997; Maddison

1997). Since here we only have a single-gene genealogy, we cannot rule out problems caused by incomplete lineage sorting of ancestral alleles.

Cyrtolophosidida is truly not monophyletic. The Cyrtolophosidida may actually be a non-monophyletic group brought together based on the combination of possibly

homoplastic characters. If this is indeed the case then we can suggest two scenarios of morphological evolution: either the members of the Cyrtolophosidida may contain pleseiomorphic characters (e.g., oral morphology), or there was convergent evolution along the two branches leading to Cyrtolophosidida I and Cyrtolophosidida II (e.g., shared outer membrane of the nuclear envelope). Further analyses of other loci are needed to test these scenarios.

2.5.9. Assignment of GenBank environmental SSU-rDNA sequences to the

Cyrtolophosididae

In this study we also placed cloned haplotypes from previously published molecular environmental diversity surveys into a phylogenetic context in the Colpodea. If we consider the close sequence similarity between HAVOmat-euk43 and LKM63 to P. alpestris, as well as the branching of these sequences inside a well-supported clade (Fig.

2.6), it is reasonable to assume that the respective organisms may indeed be tentatively assigned to the genus Pseudocyrtolophosis. Assignment to the exactly which species in

Pseudocyrtolophosis, though, we cannot say since LKM63 originates from a freshwater lake in the Netherlands (van Hannen et al. 1999) and HAVOmat-euk43 from a Hawaiian lava cave microbial mat (Brown et al., unpublished), and the morphospecies in

Pseudocyrtolophosis have likewise been isolated from a number of terrestrial environments (Foissner 1993).

Interestingly, the environmental sequence PAA10AU2004, retrieved from a

French freshwater lake (Lefèvre et al. 2007), cannot be phylogenetically assigned to any

of the sequenced genera within the family Cyrtolophosididae. Thus, it is likely that we are still far away from knowing all genera within this family.

Our analyses of the Colpodea demonstrate that increasing sampling density of known ciliate taxa in phylogenetic studies is needed in order to phylogenetically place unidentified environmental clones to known ciliate lineages as seen previously in the class Plagyopylea (Stoeck et al. 2007). By doing so, we can make predictive hypotheses of the possible morphology of the ciliates from which the clones are derived, as well as their possible metabolic and ecological roles in the environment from which they were sampled. For example, the environmental clones from previous environmental surveys placed here in the Cyrtolophosididae could be small herbivorous ciliates feeding primarily on bacteria like other taxa in the family.

2.6. Conclusions

Based on our improved observations of A. marinum it became evident that the genus Aristerostoma is a distinct taxon within the Cyrtolophosidida and probably not a junior synonym of other genera it the family, although A. minutum needs to be sampled to support this. A. marinum can be separated from other taxa in the family based on its specific infraciliature and oral structure.

Increased taxon sampling within the family Cyrtolophosididae supports an earlier analysis showing a non-monophyletic topology of the order Cyrtolophosidida.

Cyrtolophosididae again is on a separate branch from the rest of its order. Constrained analyses comparing the likelihood of the morphological placement with that of the SSU-

rDNA placement, as well as removal of the fastest evolving sites, also supports this non- monophyletic topology. However, sampling of other loci is needed to confirm these results.

In the monograph of the class Colpodea, Foissner (1993) describes only one marine species—with the remaining taxa being freshwater or terrestrial. This study, which places two potentially different species cultured from separate marine environments, points to the possibility of further Colpodea species waiting to be uncovered in future observations of marine habitats.

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Table 2.1. Taxon sampling and GenBank numbers used in this study. Newly sequenced taxa are in bold. * = sequences from environmental sampling. Taxon GB# Taxon GB# COLPODEA: Hausmanniella discoidea EU039900 Aristerostoma marinum EU264562 Ilsiella palustris EU039901 Aristerostoma sp. EU264563 Mykophagophrys terricola EU039902 Bardeliella pulchra EU039884 Notoxoma parabryophryides EU039903 Bresslaua vorax AF060453 Ottowphrya dragescoi EU039904 Bresslauides discoideus EU039885 Platyophrya-like sp. EU039905 Bryometopus atypicus EU039886 Platyophrya sp. EU039906 Bryometopus pseudochilodon EU039887 Platyophrya vorax AF060454 Bryometopus sphagni AF060455 Pseudoplatyophrya nana AF060452 Bursaria sp. 1 EU039889 Pseudocyrtolophopsis alpestris EU264564 Bursaria sp. 2 A EU039890 Rostrophrya sp. EU039907 Bursaria sp. 2 B EU039891 Sagittaria sp. EU039908 Bursaria truncatella U82204 Sorogena stoianovitchae AF300285 Chain-forming colpodid AY398684 HAVOmet-euk43* EF032797 Colpoda aspera EU039892 LKM63* AJ130861 Colpoda cucullus EU039893 PAA10AU2004* DQ244029 Colpoda inflata M97908 Colpoda steinii DQ388599 OUTGROUPS: Colpoda henneguyi EU039894 Coleps hirtus U97109 Colpoda lucida EU039895 Furgasonia blochmanni X65150 Colpoda magna EU039896 Obertrumia georgiana X65149 Colpoda minima EU039897 Orthodonella apohamatus DQ232761 Cyrtolophosis mucicola Austria EU039899 Prorodon teres X71140 Cyrtolophosis mucicola Brazil EU039898 Pseudomicrothorax dubius X65151

Table 2.2. Morphometric data of Atisterostoma marinum population Framvaren Fjord. Data are based on live observations, protargol impregnation, scanning (SEM) and transmission electron microscopy (TEM). AV=arithmetric mean; CV=coefficient of variation [%]; MA=macronucleus, max=maximum value; MI=micronucleus, min=minimum value; SE=standard error; STD=standard deviation. Morphometric min max AV STD SE CV n method character observed individuals Length [µm] 8.81 22.80 15.28 3.13 0.58 20.51 29 live 10.04 19.84 14.69 1.99 0.29 13.52 47 protargol 8.66 15.43 12.29 1.98 0.41 16.12 23 SEM

Width [µm] 3.62 10.72 7.59 1.79 0.33 23.59 29 live 5.53 10.19 7.80 1.15 0.17 14.78 47 protargol 4.98 9.02 7.14 0.93 0.19 12.96 23 SEM

number MA 1 1 n.a. n.a. n.a. n.a. 45/3 protargol/TEM

number MI 1 1 n.a. n.a. n.a. n.a. 11/4 protargol/TEM

diameter MA 2.18 4.51 3.28 0.63 0.09 19.09 45 protargol [µm]

diameter MI 1.13 1.70 1.37 0.17 0.05 12.22 11 protargol [µm]

n somatic 6 6 n.a. n.a. n.a. n.a. 23/15 SEM/protargol kineties

n oral 4 4 n.a. n.a. n.a. n.a. 5/15 SEM/protargol membranelles

n kineties of 8 8 n.a. n.a. n.a. n.a. 7 SEM paroral membrane distance MA- 4.10 9.11 6.70 1.36 0.20 20.37 45 protargol posterior end n.a. = not applicable as characters are invariable

Table 2.3. Comparative morphology of described representatives of the four genera Cyrtolophosis, Pseudocyrtolophosis, Plesiocaryon und Aristerostoma within the family Cyrtolophosididae (Ciliophora: Colpodea). DK=dikinetids; MK=monokinetids.

Figure 2.1. Light microscopy of living (a, b) and protargol impregnated (c)

Aristerostoma marinum cells. (a) The contractile vacuole (CV) and the oral structure

(OS) are visible during live observation. The photograph shows the left lateral side of the cell. (b) Cell(s) during perkinetal binary fission. (c) Due to poor impregnability (see text and Foissner 1993) protargol microphotographs are suboptimal. Nevertheless, extrusomes

(mucocsyst) are clearly visible as dark-colored dots distributed evenly in the cell’s cortex

(arrows). Also the macronucleus (MA) and the micronucleus (MI) are visible. Scale bar in all microphotographs= 5 µm

Figure 2.2. Scanning (a-c) and transmission (d, e) electron microscopy of Aristerostoma marinum. (a) left lateral - dorsal view of the cell displaying the ellipsoid shape tapering anteriorly, parts of kinety number 4 (white arrows, two dikinetids and two uniciliated kinetids are visible—see text for discussion) and four of the six kinetids (black arrows) that seem to belong to sparsely ciliated somatic kineties. Scale bar = 5 µm (b) Details of the oral apparatus with the bulge (solid white arrow), oral kinetids (black arrow) and the oral membranelle (dashed white arrow). Scale bar = 0.5 µm (c) Right lateral view with kineties 1-4 (see schematic drawing Fig. 3). Scale bar = 5 µm. (d) The black arrows point to discharged mucocysts (extrusomes) and the white arrows to mitochondria with tubular cristae. Scale bar = 2.5 µm (e) Longitudinal section of the cell showing the size, position and structure of the nuclear apparatus. The micronucleus (MIC) distinguishes from the nucleolus (NU) by its higher electron density (darker color). MIC and macronucleus

(MAC) share an outer membrane of the nuclear envelope. However, we were not able to reveal the detailed structure of the nuclear membranes (arrow). Scale bars a-c and d = 5

µm

Figure 2.3. Schematic drawing of Aristerostoma marinum (right lateral view) that combines diagnostic features as revealed by live cell imaging, protargol staining and scanning electron microscopy. For details see text.

Figure 2.4. Ecophysiological tolerance limits of Aristerostoma marinum population

Framvaren Fjord. We tested for salinity, pH, temperature and oxygen tolerance.

Figure 2.5. Schematic drawing of the idealized oral structures (paroral membranes and adoral organelles) of genera in the family Cyrtolophosididae. Note that there can be much variation among species within the genera, including the number and size of adoral organelles. In some species there may be a short oblique kinety anteriorly to the uppermost adoral organelle.

Figure 2.6. Most likely Bayesian SSU-rDNA genealogy of the class Colpodea. New sequences are in bold. Numerical support values are shown as: MP bootstrap/ML bootstrap/Bayesian posterior probability. Values <50% are shown as ‘-’. Monophyletic classes and orders are labeled with a solid line, while non-monophyletic ones are labeled with a dashed line.

CHAPTER 3

EXPANDING CHARACTER SAMPLING IN CILIATE PHYLOGENETIC

RECONSTRUCTION: MITOCHONDRIAL SSU-RDNA AS A

MOLECULAR MARKER

Micah Dunthorna, Wilhelm Foissnerb, Laura A. Katza,c

aGraduate Program in Organismic and Evolutionary Biology, University of

Massachusetts, Amherst, Massachusetts, USA bFB Organismische Biologie, Universität Salzburg, Austria cDepartment of Biological Sciences, Smith College, Northampton, Massachusetts, USA

3.1. Abstract

Ciliate molecular systematics has largely focused on increasing taxon sampling using the nuclear small subunit rDNA (nSSU-rDNA) locus. Previous nSSU-rDNA analyses have generally been congruent with the morphologically-based classification, although there is extensive non-monophyly at many levels. Nuclear protein-coding loci have been shown to be inadequate as independent phylogenetic markers because of extensive paralogy and extremely heterogenous rates of substitution. Here the mitochondrial small subunit rDNA (mtSSU-rDNA) is evaluated for deep ciliate nodes using the Colpodea as an example. Overall, well-supported nodes in the mtSSU-rDNA toplogy are congruent with well-supported nSSU-rDNA nodes within the Colpodea. The one incongruence between the loci is the placement of the Sorogenida: nSSU-rDNA nests the clade within Cyrtolophosidida II, while in the mtSSU-rDNA topology it is basal to the Cyrtolophosidida II. The placement of the Sorogenida in the concatenated topology is the same as that of mtSSU-rDNA topology. The mtSSU-rDNA and concatenated topologies are generally concordant with the classifications in that most morphological groups are supported. However, several proposed relationships are not supported by molecular data. This indicates that the morphological characters used in taxonomic circumscriptions of the Colpodea represent a mixture of ancestral and derived states. The addition of mtSSU-rDNA as a marker enables phylogenetic reconstructions of the ciliate tree of life to move from a single-locus effort to a multi-locus approach.

3.2. Introduction

Whether it is better to increase the number of sampled taxa or the number of characters to increase the accuracy of phylogenetic inference is a central debate in molecular systematics (Graybeal 1998; Hillis 1998; Rannala et al. 1998; Poe and

Swofford 1999; Hillis et al. 2003; Rokas et al. 2003; Cummings and Meyer 2005; Rokas and Carroll 2005; Hedtke et al. 2006). In general both approaches have their strengths and weaknesses, and it is advantageous to increase both when reconstructing the tree of life of any group of organisms. But this has not always been possible in all clades—such as in ciliates.

Since the discovery of ciliates (Ciliophora Doflein, 1901) by Leeuwenhoek

(Dobell 1932), our understanding of their evolutionary relationships has been improved by new techniques in visualizing morphological and developmental characters (Lynn

2008). Analyses of these morphological characters have led to several classifications and monographs for both deep and shallow relationships (Corliss 1979; Foissner 1993;

Puytorac 1994; Berger 1999; Lynn and Small 2002; Berger 2006; Foissner and Xu 2007;

Jankowski 2007; Lynn 2008). Molecular phylogenetic reconstructions to test these hypothesized deep relationships have relied primarily on expanding taxon sampling while using the nuclear small subunit ribosomal DNA (nSSU-rDNA) locus (Dunthorn and Katz

2008). Because of this single-locus approach, we do not know if the molecules are elucidating ciliate morphological evolution or just misleading us.

For deep ciliate nodes, nSSU-rDNA gene trees are concordant with many morphological hypotheses, but there are a number of discrepancies between (Lynn 2003;

Foissner et al. 2004; Dunthorn and Katz 2008; Lynn 2008). Other molecular markers

have been used for further testing of these deep ciliate nodes, but there are difficulties with them. For instance, the internally transcribed spacer region (ITS) of the rDNA locus is linked to nSSU-rDNA, and alignment is difficult because of high levels of insertions and deletions (Snoeyenbos-West et al. 2002). Nuclear protein-coding loci are problematic because of heterogeneous rates of evolution and extensive paralogy (Tourancheau et al.

1998; Israel et al. 2002; Katz et al. 2004). In contrast, there are more loci available as molecular markers for shallower ciliate nodes (Morin and Cech 1988; Sadler and Brunk

1992; van Hoek et al. 2000a; Ye and Romero 2002; Barth et al. 2006; Hori et al. 2006;

Lynn and Strüder-Kypke 2006; Przyboś et al. 2006; Chantangsi et al. 2007; Barth et al.

2008; Catania et al. 2008).

One ciliate group in which nSSU-rDNA genealogies based on increased taxon sampling have been compared to morphological hypotheses is the Colpodea Small &

Lynn, 1981. The Colpodea is diagnosed by a left kinetodesmal fiber (LKm) and unique silverline patterns (Foissner 1993; Lynn 2008). This primarily terrestrial group contains diverse oral morphologies and is potentially up to 900 million years old (Wright and

Lynn 1997; Lynn 2008). The almost 200 described species are monographed with an extensive morphological classification (Foissner 1993). Previous analyses using nSSU- rDNA have challenged some aspects of this morphologically-based classification (Lynn et al. 1999; Lasek-Nesselquist and Katz 2001; Dunthorn et al. 2008; Dunthorn et al.

2009). In light of these molecular data, modified hypotheses of morphological evolution have been proposed (Foissner and Kreutz 1998; Lynn et al. 1999; Lasek-Nesselquist and

Katz 2001; Dunthorn et al. 2008; Dunthorn et al. 2009).

Here we move ciliate systematics towards increasing character sampling by sequencing a broad sample of the Colpodea for the mitochondrial small subunit rDNA

(mtSSU-rDNA). With this character-rich locus, our approach generates an additional 823 unmasked characters for analysis that is not only from an independent locus but also one from a separate genome within ciliates. Hence, analyzing nSSU-rDNA and mtSSU- rDNA has to potential to substantially increase our power for interpreting phylogenetic relationships among ciliates and their morphological evolution.

3.3. Methods

3.3.1. Taxon sampling and terminology

Sequences were obtained from genomic DNA from earlier phylogenetic reconstructions (Dunthorn et al. 2008; Dunthorn et al. 2009) or from newly isolated material, as well as from GenBank. In total, our sampling includes 25 isolates from 24 morphospecies (Table 3.1). Of these, 20 are from the Colpodea. Exemplars from five of the seven orders within the Colpodea as recognized by Foissner (1993) are in included.

Two Paramecium species, two Tetrahymena species, and Chilodonella uncinata are included as outgroups. Initial analyses included mtSSU-rDNA hydrogenosome sequences from Armorphorea accessions in GenBank; these were excluded from the final analyses since they showed extreme rate heterogeneity compared to the rest of the sequences.

When possible, both nSSU-rDNA and mtSSU-rDNA were from the same source DNA.

Terminology follows Foissner (1993) and Lynn (2008). Classification follows Foissner

(1993), with the addition of the labeling of Cyrtolophosidida clades 1 and 2 in the tree topologies following Dunthorn et al. (2008).

3.3.2. DNA amplification and sequencing

Genomic DNA was extracted using the DNeasy Tissue kit (Qiagen, CA).

Mitochondrial SSU-rDNA was amplified with the 5’ primer (TGT GCC AGC AGC CGC

GGT AA) and the 3’ primer (CCC MTA CCR GTA CCT TGT GT) from van Hoek et al.

(2000a). Phusion polymerase (New England BioLabs, MA) was used with the following cycling conditions: 3:00 at 980; 40 cycles of 0:15 at 980, 0:15 at 670, 1:15 at 720; 10:00 extension at 720. Nuclear SSU-rDNA was amplified following Dunthorn et al. (2008).

Amplified products were cleaned with microCLEAN (The Gel Company, San

Francisco, CA), and cloned with the Zero Blunt TOPO kit (Invitrogen, Carlsbad, CA).

Positive clones were identified by PCR screening with AmpliTag Gold polymerase and vector primers (Applied Biosystems, Foster City, CA), and minipreped using Qiaprep

Spin Miniprep kit (Qiagen). Clones were sequenced with the Big Dye terminator kit

(Applied Biosystems), using vector primers. Up to eight colonies were sequenced in the forward direction, and up to five of these were also sequenced in the reverse direction.

Sequences were run on an ABI 3100 automated sequencer.

3.3.3. Genealogical analyses

Haplotypes were determined and edited from overlapping sequence reads in

SeqMan (DNAStar, Inc., Madison, WI). Vector and primer nucleotides were trimmed off.

Pairwise distances for within and among samples were calculated as uncorrected “p”

distances in PAUP* v4.0b8 (Swofford, 2002). Haplotypes were aligned using Clustal X

(Thompson et al. 1994), and further edited by eye in MacClade v4.05 (Maddison and

Maddison 2005). Ambiguously aligned regions were masked. For all datasets the GTR-I-

Γ evolutionary model was estimated using AIC in MrModeltest v2 (Nylander 2004).

Maximum parsimony (MP) analyses were carried out in PAUP* v4.0b8 (Swofford 2002), with all characters equally weighted and unordered. The TBR heuristic search option was used, running 100 random additions with MulTree option on. Maximum likelihood (ML) analyses were carried out in RAxML v7.0.4 (Stamatakis et al. 2008). Support for MP and

ML analyses came from 1000 bootstrap replicates using heuristic searches. Bayesian

Inference (BI) analyses were carried out using MrBayes v3.2.1 (Huelsenbeck and

Ronquist 2003), with support coming from posterior probability using four chains and running 10 million generations. Trees were sampled every 1000 generations. To determine if the Bayesian analyses were run long enough, output files were examined using AWTY (Nylander et al. 2008). The first 25% of sampled trees were considered burn-in trees and were discarded prior to tree reconstruction. A 50% majority rule consensus of the remaining trees was used to calculate posterior probability. Following

Holder et al. (Holder et al. 2008), this consensus tree is presented.

3.3.4. Data partitioning and congruence testing

Mitochondrial and nuclear datasets were first analyzed separately. To test if they should be combined, constrained ML analyses were carried out forcing the

Cyrtolophosididae II topologies onto the other loci. For nSSU-rDNA dataset

Platyophrya, Platyophrya-like, Rostrophrya, and Sagittaria were constrained to be

monophyletic. In the mtSSU-rDNA dataset Ottowphrya, Platyophrya, and Sorogena were constrained to be monophyletic. Resulting constrained topologies were then compared to the non-constrained ML topologies using the AU test as implemented in CONSEL v0.1j

(Shimodaira and Hasegawa 2001).

3.4. Results

3.4.1. Characteristics of gene sequences

Among sequences new here and from GenBank, the mitochondrial SSU-rDNA primers amplified fragments of variable size and G-C content (Table 3.1). For all sequences, the average number of base pairs is 1070, with a minimum of 894 in

Chilodonella uncinata and a maximum of 1196 in Colpoda magna. The average G-C content is 32.9%. Towards the five-prime end there is considerable variation in indel length (not shown).

Genetic variation in the mtSSU-rDNA locus was not found within a single isolate of the morphospecies, except in Colpoda henneguyi. The two C. henneguyi sequences are

2.69% different from each other; this same isolate had two different nSSU-rDNA sequences differing by 0.12% (Dunthorn et al. 2008). The two Cyrtolophosis mucicola isolates—one from Austria, the other from Brazil—differ by 10.053%; these two differ in their nSSU-rDNA sequences by 1.71% (Dunthorn et al. 2008). The nSSU-rDNA data from Dunthorn et al. (2008) and the mtSSU-rDNA here suggest that these two C. mucicola isolates may represent cryptic species.

3.4.2. MtSSU-rDNA analyses

The mtSSU-rDNA alignment includes 823 characters, of which 491 are parsimony-informative. The MP tree is 2334 in length. The ML tree has a -lnL of

11239.12302. There was little difference in topologies among the three methods for well- supported nodes, and the ML and BI trees are identical. Here we present the most likely

ML tree with node support from all three methods (Figure 3.1).

In all analyses the Colpodea is monophyletic, with moderate to full node support.

Support for the monophyly of the Colpodea from nSSU-rDNA has been inconsistent in previous analyses (Lynn et al. 1999; Lasek-Nesselquist and Katz 2001; Dunthorn et al.

2008). However, with the current limited taxon sampling from outgroup lineages, mtSSU-rDNA does not provide a valid test of monophyly. For those Colpodea taxa sampled here the mtSSU-rDNA topology for internal relationships is largely congruent with previous molecular phylogenetic reconstructions using nSSU-rDNA (Lynn et al.

1999; Lasek-Nesselquist and Katz 2001; Dunthorn et al. 2008; Dunthorn et al. 2009).

Mitochondrial SSU-rDNA does not support the monophyly of the

Cyrtolophosidida (Figure 3.1). The Cyrtolophosidida falls into two clades with moderately supported intervening nodes. Cyrtolophosidida I—including those taxa in the

Cyrtolophosididae—is sister to the Colpodida with no to low node support.

Cyrtolophosidida II—including the rest of the sampled Cyrtolophosidida—is sister to the rest of the Colpodea with high to full node support. This same non-monophyletic topology was found with nSSU-rDNA (Dunthorn et al. 2008; Dunthorn et al. 2009).

Using the nSSU-rDNA topology, Dunthorn et al. (2008) suggest that morphologically the

Cyrtolophosidida may represent the ancestral state within the Colpodea. These two

Cyrtolophosidida groups also differ in the number of right oral membranes: two in

Cyrtolophosidida I, and one in Cyrtolophosidida II (Dunthorn et al. 2008).

The Sorogenida is monophyletic and is sister to Cyrtolophosidida II with high to full node support (Figure 3.1). While the monophyly of the Sorogenida is also supported by nSSU-rDNA, nSSU-rDNA nests Sorogenida within the Cyrtolophosidida II clade

(Dunthorn et al. 2008; Dunthorn et al. 2009). Using the nSSU-rDNA topology, Dunthorn et al. (2008) suggest that the aerial sorocarp of Sorogena may be a complex derived character arising from within the Cyrtolophosidida II. The mtSSU-rDNA locus, while likewise suggesting a close relationship between these two groups, indicates that this complex feature arose outside the Cyrtolophosidia II. Although additional taxon sampling of previously unsequenced Cyrtolophosidia II species is needed to determine the position of these two groups, a close relationship between Sorogenida and Cyrtolophosidida II is supported in that both having brick-shaped organelles on the left side of the oral structure as well as pleurotelokinetal stomatogenesis (partial re-organization of parental oral structures during cell division) (Dunthorn et al. 2008).

In the mtSSU-rDNA topology Bryometopida and the Bursariomorphida are sister to each other with high to full node support (Figure 3.1), consistent with nSSU-rDNA topology (Dunthorn et al. 2008; Dunthorn et al. 2009). Using nSSU-rDNA topologies,

Dunthorn et al. (2008), Foissner and Kreutz (1998) and Lynn et al. (1999) note that these two groups fall sister and do share a number of morphological characters: apical oral structures, ventral clefts, ardoral organelles that are conspicuous, and cysts with emergence pores.

The Colpodida is monophyletic in the mtSSU-rDNA topology, though with no to moderate node support (Figure 3.1). Within the Colpodida, Colpoda is not monophyletic with high to full node support as the Bresslauides discoideus nested within it (Figure

3.1). The genus Colpoda was likewise not monophyletic in an earlier nSSU-rDNA analysis (Dunthorn et al. 2008). Dunthorn et al. (2008) suggest that Bresslauideus, and other groups, were split off from Colpoda because of apomorphies (e.g., semicircular right oral polykinetids) that arose from within the Colpoda clade.

3.4.3. Nuclear SSU-rDNA analyses and topology congruence

To test whether truncated taxon sampling will affect the topology of the

Colpodea, a nSSU-rDNA alignment was made with the same taxon sampling as that in the mtSSU-rDNA alignment here. This alignment includes 1631 characters, of which 438 are parsimony-informative. The two MP trees are1215 in length. The ML tree has a -lnL of 8541.68589. The MP, ML and BI trees are identical, except that in the ML tree C. magna and H. discoidea are sister to each other. Here we present the most likely ML tree with node support from all three methods of analyses (Supplementary Figure 3.1).

The nSSU-rDNA topology for the Colpodea here is the same as those previously published analyses based on greater taxon sampling (Dunthorn et al. 2008; Dunthorn et al. 2009), except for the lower node support for the clade formed by Cyrtolophosidida I and Colpodida (Supplementary Figure 3.1). The low support for this same clade in the mtSSU topology (Figure 3.1), may likewise be due to the low taxon sampling here, and may increase as more taxa are sampled for the mtSSU-rDNA.

3.4.4. Concatenated analyses

Overall, the nSSU- and the mtSSU-rDNA topologies are congruent, except in the placement of the Sorogenida. This overall congruence supports the concatenation of the two loci for a combined analysis. Furthermore, the nSSU-rDNA unconstrained topology was not significantly better than the constrained topology in the AU test (p= 0.360).

However, the mtSSU-rDNA unconstrained topology was significantly better than the constrained topology in the AU test (p= 0.046).

Given the overall congruence between the topologies, and the result of the AU test for the nSSU-rDNA dataset, a concatenated alignment of nSSU and mtSSU-rDNA was compiled. This alignment includes 2454 characters, of which 929 are parsimony- informative. The MP tree is 3573 in length. The ML tree has a -lnL of 20998.62880.

There was little difference in topologies among the three methods for well-supported nodes, and the ML and BI trees are identical. Here we present the most likely ML tree with node support from all three methods (Figure 3.2).

The nSSU-rDNA, mtSSU-rDNA, and concatenated topologies are largely congruent with each other almost all relationships. Unlike the nSSU-rDNA topology here

(Supplementary Figure 3.1) and from previous analyses (Dunthorn et al. 2008;

Dunthorn et al. 2009), the Sorogenida is sister to the Cyrtolophosidida II in both the mtSSU-rDNA and concatenated topologies (Figures 3.1, 3.2). The clade formed by

Cyrtolophosidida I and Colpodida has low node support in the concatenated topology

(Figure 3.2). This low support may likewise be due to low taxon sampling (see above).

3.5. Discussion

3.5.1. Mitochondrial SSU-rDNA as a ciliate molecular marker

Nuclear SSU-rDNA has remained the primary locus for ciliate molecular phylogenetic reconstructions since it was first sequenced by Sogin and Elwood (1986) and Lynn and Sogin (1988). Although congruent in many aspects, nuclear SSU-rDNA topologies have been used to break up or reshuffle large taxa, as well as recognize new clades (Greenwood et al. 1991; van Hoek et al. 2000b; Lynn and Strüder-Kypke 2002;

Lynn 2003; Strüder-Kypke and Lynn 2003; Affa'a et al. 2004; Gong et al. 2006; Strüder-

Kypke et al. 2006; Stoeck et al. 2007; Lynn 2008; Yi et al. 2008). Nuclear protein-coding loci have failed, so far, to provide an adequate and independent test of nSSU-rDNA topologies because of heterogeneous rates of mutation and extensive paralogy

(Tourancheau et al. 1998; Israel et al. 2002; Katz et al. 2004). The resulting reliance on just nSSU-rDNA locus to reconstruct the ciliate tree of life stands in contrast to the increasing repertoire of both low- and high-copy loci available for many other microbial and macrobial eukaryotic clades, as well as the number of loci used to reconstruct shallow ciliate relationships.

Here we show that the mtSSU-rDNA locus can provide well-supported nodes for the depths of the ciliate tree of life that were analyzed. Furthermore, most of these nodes in the individual (Figure 3.1) and concatenated (Figure 3.2) analyses are congruent with those that are well supported by the nSSU-rDNA locus in previous with previous molecular phylogenetic reconstructions using nSSU-rDNA based on greater taxon sampling (Lynn et al. 1999; Lasek-Nesselquist and Katz 2001; Dunthorn et al. 2008;

Dunthorn et al. 2009), as well as truncated taxon sampling here (Supplementary Figure

3.1).

The placement of the Sorogenida is the one well-supported incongruence between the nuclear and mitochondrial loci. In a previous analysis using nSSU-rDNA, as well as here (Supplementary Figure 3.1), the Sorogenida nests within the Cyrtolophosidida II.

Here, mtSSU-rDNA and the concatenated analyses place the Sorogenidas sister to the

Cyrtolophosidida II (Figures 3.1, 3.2). Effects from low taxon sampling may likely not explain this congruence since the taxon sampling here is the same for both nSSU- and mtSSU-rDNA. Differential rates of evolution between the loci or incomplete lineage sorting are some possible explanations for this incongruence.

The rate of substitution appears to be faster in mtSSU-rDNA than in nSSU-rDNA, such as in C. henneguyi and C. mucicola (see above). This discrepancy in rates can be explained by a number of possible factors: effective population size of the mitochondria vs. nuclei; homogenizing effects on nSSU-rDNA due to meiotic recombination, although there is debate whether the Colpodea are sexual (Foissner 1993; Dunthorn et al. 2008); and differential effects of selection among the genomes.

Further work using the mtSSU-rDNA locus is needed to test a number of taxonomic hypotheses in the Colpodea and in other ciliate groups. One approach, as done here, is to use genomic DNA previously extracted for nSSU-rDNA analyses, and reuse them to amplify mtSSU-rDNA. The other is to amplify and sequence both loci each time new genomic DNA is extracted from new isolates.

3.5.2. Morphology vs. molecules in ciliates

The potential problem that individual species trees may not necessarily be following the species tree affects all organisms (Doyle 1992; Doyle 1997; Maddison

1997). So in ciliates, when there are discrepancies between evidence from morphology and molecules, morphologists have rightly pointed out that nSSU-rDNA gene trees may not be an accurate inference of phylogeny (Agatha 2004; Foissner et al. 2004; Schmidt et al. 2007; Dunthorn et al. 2008). On the other hand, ciliate molecular systematists have pointed out that they are analyzing not only more characters for phylogenetic reconstruction, but they have also suggested alternative hypotheses, or re-interpretations, of morphological evolution given the topology of gene trees (Lynn et al. 1999; Strüder-

Kypke and Lynn 2003; Dunthorn et al. 2008; Dunthorn and Katz 2008). In general, it appears that morphologically circumscribed groups in ciliates can be based on a mixture of ancestral, derived, and convergent states, and that molecules are needed to disentangle them.

Now ciliate molecular systematists can also point to further support from analyses of mtSSU-rDNA for their interpretations and hypotheses of morphological evolution.

From the work here in the Colpodea, when there are discrepancies between morphology and nSSU-rDNA molecules, mSSU-rDNA comes down on the side of nSSU-rDNA molecules and its reinterpretation of morphological evolution. It remains to be seen if further mitochondrial analyses will likewise support re-interpretations of morphological evolution within other ciliate clades.

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3.5.3. Systematic implications

The first molecular phylogenetic reconstruction of the Colpodea by Lynn et al.

(1999) supported some aspects of the morphological classification, while others aspects were challenged. Subsequent molecular phylogenetic analyses have upheld Lynn et al.

(1999), and challenged additional aspects of the classification (Lasek-Nesselquist and

Katz 2001; Dunthorn et al. 2008; Dunthorn et al. 2009). Nevertheless, the morphologically-based taxa have remained unchanged (Puytorac 1994; Lynn and Small

2002; Adl et al. 2005; Jankowski 2007; Lynn 2008). The second-locus approach using mtSSU-rDNA is generally concordant with previous nSSU-rDNA analyses of the

Colpodea, particularly at well-supported nodes. Where there are discrepancies between morphology and molecules, this second line of evidence—from an independent locus in another genome—can be used to support a reclassification the Colpodea to reflect phylogenetic relationships.

3.6. Conclusions

Future molecular phylogenetic reconstructions of ciliate relationships can now use a two-locus approach, with both nuclear and mitochondrial SSU-rDNA. This increasing of character sampling will help bring ciliate molecular systematics up to current practices in other eukaryotic clades where the use of multiple markers is standard. Mitochondrial

SSU-rDNA topologies support previous conclusions about morphological evolution made in light of nSSU-rDNA studies.

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Table 4.1. Taxon sampling for mtSSU. GenBank numbers for new sequences are in bold. For GenBank accessions, measurements were made for only the part of the sequence that is amplified by the primers used here. mtSSU nucSSU sequence % length GC Taxon (bp) content GenBank # GenBank # Aristerostoma sp. ATCC #50986 1076 36.95 X EU264563

Bardeliella pulchra 1183 34.5 X EU039884

Bresslauides discoideus 1108 34.71 X EU039885

Bryometopus atypicus 1082 35.49 X EU039886

Bursaria muco 1116 32.77 X EU039889

Bursaria truncatella 1144 32.45 X U82204

Chilodonella unicata 894 25.5 X X

Colpoda aspera 1149 31.77 X EU039892

Colpoda cucullus 1096 31.44 X EU039893

Colpoda henneguyi 1128 31.39 X EU039894 1128 31.15 X1

Colpoda lucida 1170 30.46 X EU039895

Colpoda magna 1196 33.68 X EU039896

Cyrtolophosis mucicola Austria 1013 30.89 X EU039899

Cyrtolophosis mucicola Brazil 1015 33.33 X EU039898

Hausmaniella discoideus 1121 32.99 X EU039900

Ottowphrya dragescoi 964 31.33 X EU039904

Paramecium primaurelia 994 34.91 K01750 AF100315

Paramecium tetraurelia 992 34.98 X159172 X03772

Platyophrya sp. 1015 32.89 X EU039906

Platyophrya-like sp. 1048 34.2 X EU039905

Rostrophrya sp. 1055 34.54 X EU039907

Sagittaria sp. 1049 35.66 X EU039908

Sorogena stoianovitchae 1001 33.87 X AF300285

Tetrahymena pyriformis 1039 32.15 AF160864 M98021

Tetrahymena thermophila 1037 30.95 AF396436 X56165 1Not used in phylogenetic analyses. 2Labeled as Paramecium aurelia in GenBank.

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Figure 3.1. Mitochondrial SSU-rDNA genealogy of the Colpodea. The most likely

Bayesian tree is shown. Support values are shown as: MP bootstrap/ML bootstrap/BI posterior probability. Values <50% are shown as ‘-’. Monophyletic classes and orders are labeled with a solid line, while non-monophyletic ones labeled with a dashed line.

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Figure 3.2. Concatenated nuclear and mitochondrial SSU-rDNA genealogy of the

Colpodea. The most likely Bayesian tree is shown. Support values are shown as: MP bootstrap/ML bootstrap/BI posterior probability. Values <50% are shown as ‘-’.

Monophyletic classes and orders are labeled with a solid line, while non-monophyletic ones labeled with a dashed line.

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Supplementary Figure 3.1. Nuclear SSU-rDNA genealogy of the Colpodea. The most likely Bayesian tree is shown. Support values are shown as: MP bootstrap/ML bootstrap/BI posterior probability. Values <50% are shown as ‘-’. Monophyletic classes and orders are labeled with a solid line, while non-monophyletic ones labeled with a dashed line.

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CHAPTER 4

EXTENSIVE GENETIC DIVERSITY WITHIN HALTERIID CILIATES

Micah Dunthorna, Wilhelm Foissnerb, Laura A. Katza,c

aGraduate Program in Organismic and Evolutionary Biology, University of

Massachusetts, Amherst, Massachusetts, USA bFB Organismische Biologie, Universität Salzburg, Austria cDepartment of Biological Sciences, Smith College, Northampton, Massachusetts, USA

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4.1. Abstract

Previous analyses of two morphospecies suggest that the underlying level of genetic variation can vary among halteriid ciliates. Here sampling is increased to include more worldwide isolates with nuclear SSU-rDNA and internally-transcribed spacer (ITS) region of the SSU-rDNA locus. There is extensive genetic variation in the morphospecies

Halteria grandinella that is consistent with either a large effective population size or multiple cryptic species. This extensive genetic variation is in contrast to the little genetic variation in the close related morphospecies Meseres corlissi. Together these data point out that different ciliate morphospecies can have different underlying genetic patterns and may not be comparable in biodiversity studies.

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4.2. Introduction

Most ciliate species are circumscribed using numerous morphological, ultrastructural, and developmental characters (e.g., Berger 1999; Berger 2006; Foissner

1993; Foissner et al. 2002; Foissner et al. 2005; Lynn 2008). These morphospecies are used in biodiversity studies analyzing the potential number of ciliates species and their distributions (Finlay 2002; Finlay et al. 1996; Foissner 1999; Foissner et al. 2008).

Although morphologically circumscribed species are primarily used to understand ciliate evolution and biodiversity, there is evidence for cryptic species as some morphospecies that consist of numerous genetically distinct clades (reviewed in Foissner et al. 2008; Lynn 2008). For example, in Tetrahymena and Paramecium, cryptic species based on reproductive isolation have been described (Nanney and McCoy 1976;

Sonneborn 1937; Sonneborn 1957; Sonneborn 1975), and ecological differences within cryptic species complexes have been demonstrated (Weisse 2002). The extent of the taxonomic distribution of cryptic species in ciliates, though, is not known.

One clade that potentially has cryptic species is common, freshwater halteriid ciliates. The Halteriidae Claparède and Lachmann, 1858 have two right oral membranes

(an endoral and a reduced paroral membrane), and more than three modified somatic kineties or bristles (Agatha 2004; Foissner et al. 1991; Foissner et al. 2004; Lynn 2008;

Maeda 1986; Petz and Foissner 1992). Halteriids are model organisms used to test the debate about microbial distributions because they can be easily found and cultivated

(Foissner et al. 2008; Katz et al. 2005; Weisse et al. 2008).

One recent analysis of halteriids found evidence for potential cryptic species based on sequence divergences. Using the internally-transcribed spacer (ITS) region,

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Katz et al. (2005) examined 15 populations of the morphospecies Halteria grandinella.

ITS sequences showed three distinct clades using a 2% cutoff value. In a different morphospecies using both the small subunit rDNA (SSU-rDNA) and ITS, Weisse et al.

(2008) examined nine populations of Meseres corlissi. While their Chinese isolate is genetically different by 1% from isolates collected in Australia, Austria, and the

Dominican Republic, no consistent pattern emerged from the morphological and ecological variation among the populations.

Here, we expand on Katz et al.’s (2005) analysis of the halteriids using SSU- rDNA and ITS sequencing with more exemplars from Halteria grandinella as well as other taxa within the halteriids. We further investigate the relationships among the taxa within the Halteriidae and evaluate the possibility of cryptic species.

4.3. Materials and methods

4.3.1. Taxon sampling and identification

Seven new halteriids were isolated from six countries (Table 4.1). These new isolates were sequences for both SSU-rDNA and ITS (Table 4.2). Sequencing was also performed to complement Katz et al. (2005) so that there are both SSU-rDNA and ITS sequences for most isolates (Table 4.2). However, SSU-rDNA was not recovered from

Katz et al.’s (2005) Halteria sp. from Brazil, and hence its ITS sequence was not included in the analyses here. Halteria sequences from Katz et al. (2005) used here are:

Halteria grandinella Massachusetts (DQ241751), Halteria grandinella Colorado

(AF508759), Halteria grandinella Florida (DQ241752), Halteria grandinella

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Massachusetts (AY007444 and DQ241753), Halteria-like Dominican Republic

(DQ241757), Halteria-like Botswana (DQ241755), and Meseres corlissi Brazil

(DQ241754). Meseres sequences used here from Weisse et al. (2008) are: Austria 1

(EU339923), Austria 2 (EU399525), Austria 3 (EU339926), Austria 5 (EU399524),

Austria 6 (EU399527), Australia (EU3399528), China (EU399529), Dominican Republic

(EU399522). Species were identified using standard protocols (Foissner 1991).

4.3.2. Amplification, cloning, and sequencing

For new isolates, between 70 and 100 cells were picked with a micropipette, washed, and placed into DNA lysis buffer. Genomic DNA was extracted using the

DNeasy Tissue kit (Qiagen, CA). Genomic amplifications used Phusion polymerase following instructions (New England BioLabs, MA). Primers and cycling conditions to generate SSU-rDNA sequences followed Dunthorn et al. (2008). Primers and cycling conditions to generate ITS sequences followed Snoeyenbos-West et al. (2002).

For both Brazil samples, we also amplified SSU-rDNA through the ITS region in a single product, using the SSU-rDNA 5’ primer of Medlin et al. (1988) and the ITS 3’ primer of Snoeyenbos-West et al. (2002), with the following cycling conditions: 3:00 at

980; 36 cycles of 0:15 at 980, 0:15 at 650, and 2:30 at 720; followed by a 10:00 extension at 720. We also performed separate amplifications with an annealing temperature 700 to make sure all different copies were found. To help distinguish between natural and PCR- mediated chimeras for SSU-rDNA in the Brazil samples, we followed the recommendations of Judo et al. (1998): during amplification of the genomic DNA we

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used three times the amount of primers, and used an additional 30 seconds of extension time.

Amplified products were cloned with Zero Blunt TOPO kit (Invitrogen, CA).

Clones were screened with AmpliTaq Gold polymerase (Applied Biosystems, CA), and minipreped using Qiaprep Spin Miniprep kit (Qiagen, CA). Positive clones were sequenced with the Big Dye terminator kit (Applied Biosystems, CA) using vector and internal primers. Sequences were run on an ABI 3100 automated sequencer.

4.3.3. Genealogical analyses

Unique sequences were constructed from multiple sequence reads and edited in

SeqMan (DNAStar, Inc., Madison, WI). Vector and primer sequences were trimmed off and polymorphisms confirmed by eye. Haplotypes were scanned for potential chimeras using Chimeara (Maynard Smith 1992; Posada and Crandall 2001) as implemented in

RDP v2 (Martin and Rybicki 2000) and by eye. Chimeras were excluded from analyses.

Pairwise distances within samples were calculated as uncorrected “p” distances in

PAUP* v4.0b8 (Swofford, 2002). SSU-rDNA and ITS haplotypes were aligned in Clustal

W (Thompson et al. 1994) as implemented in DNAStar. Alignments were further edited by eye in MacClade v4.05 (Maddison and Maddison 2005). Ambiguously aligned regions were conservatively masked, and remaining gaps were treated as missing data.

Each locus was analyzed separately, and then concatenated. The GTR+I+G evolutionary model was estimated for each alignment using AIC in MrModeltest v2

(Nylander 2004). Maximum parsimony (MP) analyses were carried out in PAUP* v4.0b8

(Swofford 2002) running 100 replicates with MulTrees on. Maximum likelihood (ML)

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analyses were carried out in RAxML (Stamatakis et al. 2008). Support for MP and ML analyses came from 1000 bootstrap replicates using heuristic searches. Bayesian

Inference (BI) were carried out in MrBayes v3.1 (Huelsenbeck and Ronquist 2003) with support coming from posterior probability using four chains and running 5 million generations and sampling every 100. The first 25% of sampled trees were considered burn-in trees and were discarded prior to tree reconstruction.

Initial analyses including exemplars from all Spirotrichea clades (the potential outgroups) showed halteriid SSU-rDNA sequences to be monophyletic and nesting within stichotrich ciliates (data not shown). Furthermore, these broad initial analyses showed Meseres sequences basal to all other halteriid ciliates (data not shown). Only halteriida sequences were used in the analyses below with Meseres sequences rooting the

SSU-rDNA, ITS, and concatenated trees.

4.4. Results

4.4.1. Intra-isolate pairwise distances

The SSU-rDNA locus had no intra-isolate variation for any but the two new

Brazilian isolates (Table 4.1). The first Brazilian isolate contained two different sequences with a pairwise distance of 1.82% (Brazil 1.1, Brazil 1.2). The second

Brazilian isolate also had two different sequences with a distance of 1.67% (Brazil 2.1,

Brazil 2.2). Within the amplifications of the second Brazilian isolate we also found numerous chimeric sequences (data not shown). Each chimera sequence was found only once in each of the twelve separate amplifications, while the two main sequences where

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found in all amplifications. With an increase in primer concentration and extension time, these chimeras were almost eliminated. Hence, we consider these chimeras to be PCR- mediated and not included in the analyses.

The ITS locus had no intra-isolate variation for all but the new Brazilian isolates and the Australian isolate (Table 4.1). The first Brazilian isolate contained two different sequences with a pairwise distance of 2.9%. The second Brazilian isolate had two different sequences with a distance of 3.56%. The Australian isolate also had two different sequences with a distance of 0.19%. In all preliminary analyses these two sequences formed a clade; therefore, only one is presented here.

4.4.2. Genealogies

The SSU-rDNA alignment includes 1705 characters, of which 56 are parsimony- informative. The 80 MP trees are 168 in length. The ML tree has a –lnL of 3491.62804.

The BI tree has –lnL of 3487.65229. The ITS alignment for the Halteriidae includes 520 characters, of which 73 are parsimony-informative. The 45 MP trees are 156 in length.

The ML tree has a –lnL of 1631.19274. The BI tree has a –lnL of 1630.86044. The concatenated SSU-rDNA and ITS alignment includes 2225 characters, of which 129 are parsimoniously informative. The two MP trees are 330 in length. The ML tree has a –lnL of 5246.37838. The BI tree has a –lnL of 3255.32580. The most likely BI tree from the concatenated analysis is shown and discussed below, with node support from MP bootstrap, ML bootstrap, and BI posterior probabilities (Fig. 4.1). Individual gene trees are in the supplement (Supplements 4.1-4.2). Overall, well-supported nodes in the SSU- rDNA, ITS, and combined analyses are congruent.

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All isolates of M. corlissi are monophyletic with full node support (Fig. 4.1). The

Brazilian M. corlissi isolate characterized here is almost identical to the other M. corlissi characterized by Weisse et al. (2008). The minimal overall genetic variation found by

Weisse et al. (2008) in the morphospecies M. corlissi is supported here.

In the concatenated tree (Fig. 4.1), the Botswana Halteria-like isolates forms clade with the Dominican Republic Halteria-like isolate. This clade is in turn sister to the

Halteria grandinella morphospecies with no to low node support. In the SSU-rDNA tree

(Supplement 4.1), the Botswana isolate is sister to all other Halteria-like isolates with no node support. In the ITS tree (Supplement 4.2), this sequence is sister to the clade that includes some isolates of Halteria grandinella with moderate ML and BI node support.

The Dominican Republic isolate is sister to one of the sequences from the Brazil isolates with high to full node support (Fig. 4.1).

The genomic DNA from the two Brazilian populations newly isolated for this study each contained two different SSU-rDNA and ITS sequences. For both of these, cells were taken from a communal culture and may have contained more than one morphospecies. One sequence, Brazil 2.2, was almost exactly like that of the Dominican

Republic isolate. The other SSU-rDNA/ITS sequence (Brazil 2.1) nested within the core

Halteria grandinella clade with no node support (Figs. 4.1). The other Brazil isolate’s two sequences (Brazil 1.1 and Brazil 1.2) formed a clade in the SSU-rDNA and concatenated tree with no to moderate node support (Fig. 4.1, Supplement 4.1), but are paraphyletic in the ITS tree with no node support (Supplement 4.2).

The Halteria grandinella morphospecies isolated by Katz et al. (2005) and those from GenBank—Colorado, Ecuador, Florida, and Massachusetts—form a clade with the

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new Peru isolate in all analyses with high to full node support (Fig. 4.1). Some of the other morphologically similar Halteria grandinella specimens isolated for this study—

Africa, Australia, China, and Venezuela—formed another clade in all analyses with variable support (Fig. 4.1). In the ITS and concatenated tree these two clades formed a large clade with each other (and the Brazil 2.1 sequence) with low to no node support

(Fig. 4.1, Supplement 4.2).

4.5. Discussion

4.5.1. Genetic variation underlying morphospecies

In the SSU-rDNA, ITS, and concatenated analyses two distinct clades were uncovered with full support from all methods: 1) the morphospecies Meseres corlissi sequences; and 2) the other with Halteria sequences, which includes the morphospecies

Halteria grandinella and Halteria-like species (Fig. 4.1). These two clades are strikingly different. There is minimal genetic variation within Meseres corlissi. In other words, if you go out and collect Meseres corlissi from various locations you seem to get only one genetic entity. In contrast, there is much genetic variation within Halteria grandinella.

Two possible reasons may underlie the extensive genetic variation within

Halteria grandinella morphospecies. The different genetic subclades may represent cryptic sexual species as sex is known in Halteria grandinella (Agatha and Foissner

2009). Reproductive isolation experiments can be conducted to support this hypothesis.

Alternatively, the different genetic subclades may be the result of a very large effective population size. Halteria grandinella is easily found in most freshwater environments

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throughout the world, so census populations are large. Although the data here is consistent with a large population, we see no evidence of recombination among sequences as would be expected from a large interbreeding population. Although large population sizes are often assumed for ciliates (Finlay 2002), molecular support for large effective population sizes is conflicting (Catania et al. 2008; Katz et al. 2006; Lynch and

Conery 2003; Snoke et al. 2006). Further molecular investigations using protein-coding loci are needed to test for effective population size.

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4.6. References

Agatha, S. 2004. A cladistic approach for the classification of oligotrichid ciliates (Ciliophora: Spirotricha). Acta Protozool. 43:201-217.

Agatha, S., and W. Foissner. 2009. Conugation in the spirotrich ciliate Halteria grandinella (Müller, 1773) Dujardin, 1841 (Protozoa, Ciliophora) and its phylogenetic implications. Eur. J. Protistol. 45:51-63.

Berger, H. 1999. Monograph of the Oxytrichidae (Ciliophora, Hypotrichia). Monographiae Biol. 78:i-xii, 1-1080.

Berger, H. 2006. Monograph of the Urostyloidea (Ciliophora, Hypotrichia). Monographiae Biol. 85:i-xv, 1-1303.

Catania, F., F. Wurmser, A. A. Potekhin, E. Przyboś, and M. Lynch. 2008. Genetic diversity in the Paramecium aurelia species complex. Mol. Bio. Evol. 26:421- 431.

Dunthorn, M., W. Foissner, and L. A. Katz. 2008. Molecular phylogenetic analysis of class Colpodea (phylum Ciliophora) using broad taxon sampling. Mol. Phylogenet. Evol. 48:316-327.

Finlay, B. J. 2002. Global dispersal of free-living microbial eukaryote species. Science 296:1061-1063.

Finlay, B. J., G. F. Esteban, and T. Fenchel. 1996. Global diversity and body size. Nature 383:132-133.

Foissner, W. 1991. Basic light and scanning electron microscopic methods for taxonomic studies of ciliated protozoa. Europ. J. Protistol. 27:313-330.

Foissner, W. 1993. Colpodea (Ciliophora). Protozoenfauna 4/1:i-x, 1-798.

Foissner, W. 1999. Protist diversity: estimates of the near-imponderable. Protist 150:363-368.

Foissner, W., H. Berger, K. Xu, and S. Zechmeister-Boltenstern. 2005. A huge, undescribed soil ciliate (Protozoa: Ciliophora) diversity in natural forest stands of Central Europe. Biodivers. Conserv. 14:617-701.

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Foissner, W., H. Blatterer, H. Berger, and F. Kohmann. 1991. Taxonomische und ökologische Revision der Ciliaten des Saprobiensystems—Band I: Cyrtophorida, Oligotrichida, Hypotrichia, Colpodea. Informationsberichte Bayer. Landesmat für Wasserwirtschaft, München

Foissner, W., A. Chao, and L. A. Katz. 2008. Diversity and geographic distribution of ciliates (Protista: Ciliophora). Biodivers. Conserv. 17:345-363.

Huelsenbeck, J. P., and F. R. Ronquist. 2003. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19:1572-1574.

Judo, M. S. B., A. B. Wedel, and C. Wilson. 1998. Stimulation and suppression of PCR- mediated recombination. Nucleic Acids Res. 26:1819-1825.

Katz, L. A., G. B. McManus, O. L. O. Snoeyenbos-West, A. Griffin, K. Pirog, B. Costas, and W. Foissner. 2005. Reframing the 'Everything is everywhere' debate: Evidence for high gene flow and diversity in ciliate morphospecies. Aquat. Microb. Ecol. 41:55-65.

Katz, L. A., O. Snoeyenbos-West, and F. P. Doerder. 2006. Patterns of protein evolution in Tetrahymena thermophila: implications for estimates of effective population size. Mol. Biol. Evol. 23:608-614.

Lynch, M., and J. S. Conery. 2003. The origins of genome complexity. Science 302:1401-1404.

Lynn, D. H. 2008. The ciliated protozoa: characterization, classification, and guide to the literature, 3rd edition. Springer, Dordrecht.

Maddison, W. P., and D. R. Maddison. 2005. MacClade v. 4.0.8. Sinauer Assoc.

Maeda, M. 1986. An illustrated guide to the species of the families Halteriidae and Strobilidiidae (Oligotrichida, Ciliophora), free swimming protozoa common in the aquatic environment

Martin, D., and E. Rybicki. 2000. RDP: detection of recombination amongst aligned sequences. Bioinformatics 16:562-563.

Maynard Smith, J. 1992. Analyzing the mosaic structure of genes. J. Mol. Evol. 34:126- 129.

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Medlin, L., H. J. Elwood, S. Stickel, and M. L. Sogin. 1988. The characterization of enzymatically amplified eukaryotes 16S-like ribosomal RNA coding regions. Gene 71:491-500.

Nanney, D. L., and J. W. McCoy. 1976. Characterization of the species of the Tetrahymena pyriformis complex. Trans. Am. Microsc. Soc. 95:664-682.

Nylander, J. A. 2004. MrModeltest v2. Distrubuted by the author. Evolutionary Biology Center, Uppsala University, Upsalla.

Petz, W., and W. Foissner. 1992. Morphology and morphogenesis of Strobilidium caudatum (Fromentel), Meseres corlissi n. sp., Halteria grandinella (Muller), and Strombidium rehwaldi n. sp., and a proposed phylogenetic system for oligotrich ciliates (Protozoa, Ciliophora). J. Protozool. 39:159-176.

Posada, D., and K. A. Crandall. 2001. Evaluation of methods for detecting recombination from DNA sequences: computer simulation. Proc. Natl. Acad. Sci. USA 98:13757-13762.

Snoke, M. S., T. U. Berendonk, D. Barth, and M. Lynch. 2006. Large global effective population sizes in Paramecium. Mol. Biol. Evol. 23:2474-2479.

Sonneborn, T. M. 1937. Sex, sex inheritance and sex determination in Paramecium aurelia. Proc. Natl. Acad. Sci. 23:378-383.

Sonneborn, T. M. 1957. Breeding systems, reproductive methods, and species problems in protozoa. Pp. 155-324 in E. Mayr, ed. The species problem. American Association for the Advancement of Science, Washington D. C.

Sonneborn, T. M. 1975. The Paramecium aurelia complex of fourteen sibiling species. Trans. Am. Microsc. Soc. 94:155-178.

Stamatakis, A., P. Hoover, and J. Rougemont. 2008. A fast boostrap algorithm for the RAxML web-servers. Syst. Biol. 57:758-771.

Swofford, D. 2002. PAUP*. Phylogenetic Analysis Using Parsimony (*and Other Methods). Sinauer Assoc, Sunderland MA.

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Weisse, T. 2002. The significance of inter- and intraspecific variation in bacterivorous and herbivorous protists. Antonie Van Leeuwenhoek 81:327-341.

Weisse, T., M. C. Strüder-Kypke, H. Berger, and W. Foissner. 2008. Genetic, morphological, and ecological diversity of spatially separated clones of Meseres corlissi Petz & Foissner, 1992 (Ciliophora, Spirotrichea). J. Eukaryot. Microbiol. 55:257-270.

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Table 4.1. Isolates newly collected for this study. Designation Taxon Place of collection, latitude/longitude in analyses Halteria grandinella Foissner 1: Venezuela Venezuela Halteria grandinella Foissner 2: Brazil, Pantanal (Meseres site) Brazil 1 Halteria grandinella Foissner 3: Peru, Late Titicaca Peru Halteria grandinella Foissner 4: China, Pearl River Floodplain China Halteria grandinella Foissner 5: Africa, Kruger National Park Africa Halteria grandinella Foissner 6: Brazil, Pantanal (Site 1 of Maria & Birgit) Brazil 2 Halteria grandinella Foissner 7: Australia, site 1/2006 Australia

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Table 4.2. Newly sampled loci. Sequencing of clone was performed with just the 5' primer (partial) or with the 3' and internal primers are well (full). Tbd = to be determined.

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Figure 4.1. Concatenated SSU-rDNA/ITS genealogy of the halteriids. The most likely

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Supplementary Figure 4.1. SSU-rDNA genealogy of the halteriids. The most likely

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Supplementary Figure 4.2. ITS genealogy of the halteriids. The most likely Bayesian tree is shown. Support values are shown as: MP bootstrap/ML bootstrap/BI posterior probability. Values <50% are shown as ‘-’. Monophyletic classes and orders are labeled with a solid line, while non-monophyletic ones labeled with a dashed line.

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CHAPTER 5

RICHNESS OF MORPHOLOGICAL HYPOTHESES IN CILIATE

SYSTEMATICS ALLOWS FOR DETAILED ASSESSMENT OF HOMOLOGY

AND COMPARISONS WITHGENE TREES

Micah Dunthorna and Laura A. Katza,b

aGraduate Program in Organismic and Evolutionary Biology, University of

Massachusetts, Amherst, Massachusetts, 01003, USA bDepartment of Biological Sciences, Smith Collage, College Road, Northampton,

Massachusetts, 01063, USA

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5.1. Abstract

Morphological investigations are central to ciliate systematics. Morphology has provided most species delimitations as well as almost all hypotheses on the ciliate tree of life.

Moreover, emerging analyses of molecular markers are generally concordant with morphology-based ciliate taxonomies. Despite the richness of morphology-based hypotheses, there are challenges to ciliate morphological systematics that include the decreasing numbers of trained morphologists and the difficulty in establishing homology for some morphological traits. There are also open questions about ciliate morphology, such as the cause of morphological stasis in cryptic species, and the contrasting pattern of considerable morphological variation with little underlying genetic variation.

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5.2. Introduction

Ciliates are unique among microbial groups in that their diverse morphology, abundance and relatively large sizes have enabled the creation of a comprehensive morphology-based taxonomy. Analyses of morphological characters, first gathered using light microscopy and more recently in analyses of electron micrographs, have led to detailed hypotheses on the relationships among ciliates that extend across taxonomic levels. Hence, those of us working on the systematics of ciliates find ourselves in the enviable position of having numerous hypotheses that can be assessed through both reexamination of morphology and characterization of molecular characters. Here, we describe the strengths of morphological approaches to ciliate taxonomy, the challenges to these types of analyses, the concordance between morphological and molecular characters, and the nature of some of the open questions in ciliate systematics.

5.3. Strengths of Morphology

5.3.1. Species delimitations

The diverse morphology among ciliates has allowed for many in-depth studies that have defined the limits of ciliate species. In general, the morphological species concept is the standard for ciliates (e.g., Foissner et al. 2002), although species have been named using other methods (e.g., Foissner and Berger 1999; Nanney and McCoy 1976; e.g., Sonneborn 1975). These morphological investigations provide us with an estimate of the minimal number of extant ciliate species. Although it is argued that the number is around 3000 by ecologists (Finlay 2002; Finlay et al. 1996), the actual number may be

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“near- imponderable” (Foissner 1999). Estimates of ciliate species numbers require highly trained taxonomists exploring new environments and different parts of the Earth: the more they look, the more they find (e.g., Berger 1999; Berger 2006; Berger et al.

2006; Foissner 1994; Foissner 1995; Foissner 1997a; Foissner 1998; Foissner 2003;

Foissner 2005a; Foissner et al. 2002; Foissner et al. 2005a; Foissner and Stoeck 2006;

Foissner et al. 2003; Foissner and Xu 2007; Foissner et al. 2005c; Kim et al. 2007; Lin et al. 2007; Ma et al. 2006; Petz et al. 1995). Molecular investigations using environmental sampling of SSU-rDNA haplotypes also points to such a high number (Doherty et al.

2007; Stoeck et al. 2006).Hence, there are likely many more ciliates that have yet to be discovered, maybe even up to 30,000 to 40,000 (Chao et al. 2006; Foissner 1997b;

Foissner 1999; Foissner et al. 2008).

5.3.2. Ciliate tree of life

Analyses of morphological characters, including somatic and oral ciliature, and ontogenesis have generated almost all hypotheses on the topology of the ciliate tree of life for the most inclusive clades (Corliss 1979; Lynn and Small 1997; Lynn and Small

2002; Puytorac 1994; Small and Lynn 1981; Small and Lynn 1985). Recently, morphological depictions, along with supporting molecular evidence, have divided ciliates into two subphyla—the Postciliodematophora and Intramacronucleata—and eleven classes (Adl et al. 2005; Lynn 2003). Detailed morphological hypotheses have also generated almost all hypotheses of relationships within these eleven classes (e.g.,

Berger 1999; Berger 2006; Foissner 1993; Foissner and Xu 2007; Lynn and Small 2002;

Matthes et al. 1988).

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5.4. Challenges to Morphological analyses

5.4.1. Decline in number of trained taxonomists

Like in all eukaryotic clades (Lee 2000; Wheeler 2008), one principal impediment to our understanding of ciliate diversity is the lack of trained morphological taxonomists.

As much of science in the past century shifted to a focus on model organisms, fewer and fewer students received training in collection, identification and analysis of diverse lineages, particularly microorganisms. This problem is particularly acute today as there is increasing interest in microbial diversity on Earth but few professors positioned to train students in microbial morphological taxonomy.

5.4.2. Number of characters

While the number of characters needed for phylogenetic analyses is debated

(Gatesy et al. 2007; Rokas et al. 2003), morphological characters are limited, particularly when compared to molecular characters (Givnish and Sytsma 1997; Hillis and Wiens

2000; Scotland et al. 2003). In ciliates this lack of numerous unambiguous morphological characters remains problematic, particularly when compared to most macrobes (e.g., Doyle and Endress 2000; Giribet and Wheeler 2002). In light of this, we agree with Scotland et al. (2003) that it may be more critical for morphological studies to investigate fewer characters in depth, such as in the studies of the cysts of Meseres corlissi (Foissner 2005b; Foissner et al. 2005b; Foissner and Pichler 2006; Foissner et al.

2006).

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5.4.3. Homology assessment

Likewise, homology assessment of morphological characters can be difficult in all eukaryotic clades (Scotland et al. 2003), and it is not surprising that this problem is amplified in microbial groups. We agree that homology is equated with synapomorphy

(de Pinna 1991; Patterson 1982; Stevens 1984) and hence, establishing homology is essential for inferring evolutionary relationships. Establishing homology is a two-step process. In a primary homology assessment, similarity among characters is initially established and shared ancestry is hypothesized. In a secondary homology assessment, the primary assessment is tested via congruence with other morphological or molecular characters (de Pinna 1991). Hence, inferring robust phylogenies based requires independent data sets and reassessment of primary homology statements.

While primary homology statements in ciliates can be relatively straight forward, establishing secondary homology statements is problematic because of issues in executing congruence tests. First, cladistic analyses using morphological characters in ciliates are rare and often deal with few taxa (e.g., Agatha 2004; Agatha and Strüder-

Kypke 2007; Berger and Foissner 1997; Foissner et al. 2007; Puytorac et al. 1994). As a result, most primary homology statements just have not been tested. Like in all systematic analyses, there is a difficulty in these few ciliate examples of how many taxa—both ingroup and outgroup—need to be sampled and coded for; the paper by

Foisser et al. (2007) stands out in increasing outgroups for the problem of placing the

Halteriids.

Second, the question of the level of generality of the homology of many ciliate morphological characters remains unresolved because most molecular estimates of ciliate

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relationships rely on single locus, SSU-rDNA (Dunthorn et al. 2008; Hewitt et al. 2003;

Schmidt et al. 2007a; Schmidt et al. 2007b; Shin et al. 2000; Snoeyenbos-West et al.

2004; Snoeyenbos-West et al. 2002; Strüder-Kypke et al. 2007; Strüder-Kypke et al.

2006; Williams and Clamp 2007). With the well-known gene tree versus species tree problem (Doyle 1997; Maddison 1997), we do not know yet if the SSU-rDNA locus accurately reflects species phylogeny. For example, the homology of Halteriid oral membranes with other spirotrichs remains to be satisfactorily answered, although there are numerous hypotheses (explicit or implicitly implied) and molecular tests (Agatha

2004; Agatha and Strüder-Kypke 2007; Foissner et al. 2004; Foissner et al. 2007; Petz and Foissner 1992; Strüder-Kypke and Lynn 2003; Szabó 1935). Development of additional molecular markers is essential for robustly testing homology.

5.5. Concordance with molecular hypotheses

Hypotheses of the ciliate tree of life are generally congruent with gene trees. For example, the most inclusive clades proposed—ranked at the class level—have, for the most part, either been supported or at least not refuted by molecules (Lynn 2003).

Molecules also generally support less inclusive ciliate clades. For example, much of

Foissner’s (1993) morphological classification of the class Colpodea is largely congruent with SSU-rDNA gene trees (Dunthorn et al. 2008; Lasek-Nesselquist and Katz 2001;

Lynn et al. 1999).

One aspect of the incongruence between the morphological hypotheses and the

SSU-rDNA gene trees in the Colpodea centers upon paraphyletic groups; e.g., the

Sorogenida nesting within part of the Cyrtolophosidida, the Bursariomorphida nesting

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within the Bryometopida, and the Grossglockneriida nesting within the Colpodida

(Dunthorn et al. 2008). In these three cases of paraphyly there are a number of morphological characters that unite the respective groups within the gene trees (Dunthorn et al. 2008). Another aspect of the incongruence is the challenge that plesiomorphic characters pose when trying to uncover evolutionary relationships, where the ancestral condition of the group remains in some taxa, causing them to be grouped together. An example of this is in the possibility of the Cyrtolophosidida being polyphyletic (Dunthorn et al. 2008).

5.6. Open Questions

5.6.1. Lack of morphological variation when there is genetic diversity

While morphology provides us with the minimal number of extant species, there are undoubtedly many more. Like in other eukaryotic clades (Mayr 1963; Pfenninger and

Schwenk 2007), cryptic species are well-known in ciliates (Sonneborn 1937; Sonneborn

1957). Underlying these ciliate cryptic species there can be both a high genetic diversity as well as ecological variation (Foissner et al. 2008; Katz et al. 2005; Nanney et al. 1998;

Simon et al. 2008; Weisse and Lettner 2002; Weisse and Rammer 2006).

Two main reasons why there are cryptic species have been postulated: the species may be nascent, with little time to acquire morphological difference; or the conserved morphology of the species may be of selective value (Mayr 1976). This second reason is generally accepted for ciliates (Nanney 1977; Nanney 1982; Nanney 1999; Nanney et al.

1998). This selective reason is supported by the hypothesis that cryptic ciliates may be

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ancient clades, although the actual age is debated (Nanney 1977; Nanney 1982; Nanney

1999; Van Bell 1985)—but the data on which these ages are set are weak to nonexistent.

There have been no tests, though, of the selective value of keeping the same morphology among cryptic ciliate species. Equally plausible is that selection is not occurring at all on morphology and that morphology remains in stasis for other reasons such as constraints or canalization. Alternatively, the prevalence of cryptic species of ciliates may be due to disparate rates of morphological and molecular evolution enabled by the dual nature of ciliate genomes. The separation of genome function between the unexpressed germline micronucleus and the expressed somatic macronucleus changes the dynamics of molecular evolution in ciliates as compared to other eukaryotes. Based on analyses of multiple molecular markers in diverse ciliate, the dual nature of genome evolution has been shown to be related to elevated rates of protein evolution in this lineage (McGrath et al. 2006; Zufall et al. 2006). This elevated rate of molecular evolution, coupled with the prevalence of epigenetics in development (McGrath et al.

2006), may contribute to the generation of cryptic species.

5.6.2. Lack of genetic diversity when there is morphological variation

In contrast to cryptic species, there are also cases in ciliates in which there is only limited genetic variation, at least as measured by SSU-rDNA divergence, in light of considerable morphological variation. This phenomenon is best seen in comparisons among SSU-rDNA gene trees of various clades in the class Spriotrichea. Morphological and molecular changes are relatively concordant among members of the choreotrichs, oligotrichs; in contrast, there is considerable discordance and very short SSU-rDNA

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branches among stichotrich taxa (Agatha and Strüder-Kypke 2007; Snoeyenbos-West et al. 2002; Strüder-Kypke and Lynn 2003). Intriguingly, it is only within the stichotrichs that we have evidence of gene scrambling, a process whereby coding domains are reshuffled in ciliate micronuclei (Ardell et al. 2003; Greslin et al. 1989; Prescott 1992).

We hypothesize that this type of heritable scrambling can cause instant, or at least rapid, speciation as extensive gene scrambling will disrupt pairing of homologous chromosomes. Under this scenario, accumulation of scrambled genes within populations can lead to a barrier to gene exchange with other populations of the same species.

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CHAPTER 6

ANCIENT ASEXUAL LINEAGES OF MICROBIAL EUKARYOTES THAT

REGAINED SEX?

Micah Dunthorna and Laura A. Katza,b

aGraduate Program in Organismic and Evolutionary Biology, University of

Massachusetts Amherst, USA bDepartment of Biological Sciences, Smith College, USA

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6.1. Abstract

Sex in ciliates occurs through mutual exchange of haploid nuclei during conjugation, a process that is decoupled from cell division. Among ciliates in the clade Colpodea only

Bursaria truncatella is known to have sex. Here we review the evidence for and against sexuality in the rest of the Colpodea. We discuss expectations of sexuality in light of the ancient age of the Colpodea and the problem of reversing the loss of sex in B. truncatella.

Based on these arguments, we suggest that many, if not all, of the Colpodea may be sexual. These expectations and arguments, though, are derived from theories and observations from macrobes, and may not apply to microbial eukaryotes such as ciliates.

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6.2. Introduction

Sex (amphimixis) in the ancestor of extant eukaryotes was likely facultative

(Dacks and Roger 1999). While most taxa have remained sexual, asexual lineages are found scattered throughout the eukaryotic tree of life, primarily at the tips (Bell 1982).

The pattern where most eukaryotes are sexual has been explained by theories on the advantages of the maintenance of sex; i.e., the Red Queen, Muller’s ratchet, and others

(Arkhipova and Meselson 2004; Bell 1982; Bell 1988; Burt 2000; Fischer 1930;

Hamilton 2001; Kondrashov 1982; Kondrashov 1993; Lynch et al. 1993; Maynard Smith

1978; Muller 1964; West et al. 1999; Williams 1975).

Ciliates—a clade of microbial eukaryotes—have remained facultatively sexual.

Sex in ciliates occurs during conjugation, where haploid nuclei are mutually exchanged between complementary cells (Dini and Nyberg 1993; Lynn 2008; Sonneborn 1957).

These nuclei fuse to make a zygotic nucleus that mitotically divides to give rise to a

“germline” micronucleus and a “somatic” macronucleus. The micronucleus can divide meiotically to produce the haploid nuclei that take part in conjugation (Lynn 2008;

Raikov 1996). Sex is assumed to occur in almost all ciliate clades, although details and direct observations for most species is lacking. There are known derived asexual strains that have lost their micronuclei and are thus unable to conjugate (Bell 1988; Lynn 2008).

The Colpodea—one of eleven major ciliate lineages—consists of about 200 species with similar somatic but diverse oral morphologies (Foissner 1993; Lynn 2008).

Colpodeans can be found in numerous habitats, some are fungivores, and at least one species has a multicellular life stage (Foissner 1993). Much is known about colpodeans and their evolution through morphological and molecular analyses (Dunthorn et al. 2009;

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Dunthorn et al. 2008; Foissner 1985; Foissner 1993; Lasek-Nesselquist and Katz 2001;

Lynn 1976; Lynn 2008; Lynn et al. 1999). However, there is a lack of consensus on a fundamental aspect of their biology—their sexuality. Although all known species have micronuclei, conjugation has been observed only in Bursaria truncatella (Foissner 1993;

Poljansky 1934). The extent of sexuality in the rest of the colpodeans is debated: Foissner

(1993) proposes that they are asexual, while Dunthorn et al. (2008) suggest that they are covertly sexual. It may not be surprising that we know little about sexuality in colpodean ciliates. Even in known sexual ciliate species conjugation is not always easy to induce in the laboratory (Sonneborn 1957). Ciliates also lack sex-specific morphologies or organs, so you cannot look for morphologically different sexes and at least assume they are sexual.

Here we review the evidence for and against sexuality in colpodeans. We also review two reasons why we suspect the colpodeans to be sexual: the problems of ancient asexuality and reversing the loss of sex.

6.3. Empirical evidence

While sex is well established in B. truncatella there is conflicting evidence for or against sex in the rest of the colpodeans. Foissner (1993) proposes that they are asexual because conjugation has not been observed even though numerous species have been examined over many years. Likewise, darwinulid ostracods were long thought to be asexual because males were never seen until they were found after a century of searching (Smith

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et al. 2006). Asexuality of the colpodeans may represent a similar situation to these ostracods: given enough time and requisite conditions, conjugation might be observed.

Two molecular studies of the colpodeans provide conflicting evidence. Bowers et al. (1998) found that large subunit rDNA restriction digests of three species in the clade

Colpoda showed limited to no genetic variation within local populations. They suggest that this data is consistent with asexuality. However, these molecular data are also consistent with either sexuality (leading to homogenization of variation) or infrequent sex

(leading to clonality). This study also was not designed to test directly for the presence or absence of sex.

In another molecular study not explicitly designed to test for sex, Dunthorn et al.

(2008) found little to no variation among small subunit rDNA sequences from a broad sample of colpodeans. They suggest that this pattern is consistent with sexuality because asexuality is hypothesized to lead to high intra-isolate allelic divergence because of the absence of recombination (i.e., the Meselson effect; Birky 1996; Mark Welch and

Meselson 2000; Normark et al. 2003). However, testing for the Meselson effect is problematic, as it can be masked or mimicked by a number of processes such as non- meiotic recombination or rare sex (Ceplitis 2003; Normark et al. 2003; Omilian et al.

2006). Also ciliates, because of their sometimes non-canonical genetics and high levels of paralogy (Katz et al. 2004; Nanney 1980; Snoeyenbos-West et al. 2002), pose additional challenges to interpreting molecular signatures. Future studies could try to test for or against sexuality by looking at other genomic signatures; e.g., the degree of decay of sex- related loci, increased rate of deleterious mutations, number of retrotransposons, etc.

(Normark et al. 2003). However, ciliates might pose challenges to these tests are well.

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6.4. Are colpodeans ancient asexuals?

If colpodeans are asexual (except B. truncatella) they would be the oldest putative ancient asexual lineage. Molecular clock estimates place their age up to 900 million years old (Wright and Lynn 1997). There are also fossils in 93 million year old amber (Martín-

González et al. 2008; Schmidt et al. 2001; Schönborn et al. 1999). Colpodeans are thus at least as old, or perhaps even an order of magnitude older, than the putatively ancient asexual bdelloid rotifers that date to about 130 million years old (Mark Welch et al.

2008).

While asexuals are known at the tips of the eukaryotic tree of life, ancient asexuals are generally thought to be unlikely because asexuality is believed to lead to rapid extinction (Lynch et al. 1993; Maynard Smith 1978; Normark et al. 2003)—but see

Schwander and Crespi (2009) for an alternative view. For example, without sex eukaryotic lineages may be more susceptible to increased mutational load and retrotransposons, may not be able to adapt to a changing environment, or may not be able to escape predators and parasites over evolutionary time compared to those lineages that remain sexual (Arkhipova and Meselson 2004; Bell 1982; Burt 2000; Hamilton 2001;

Kondrashov 1993; Maynard Smith 1978). Most claims of ancient asexuals have not been supported (Judson and Normark 1996; Normark et al. 2003), except possibly the bdelloid rotifers (Arkhipova and Meselson 2000; Mark Welch et al. 2004a; Mark Welch et al.

2008; Mark Welch and Meselson 2000; Mark Welch et al. 2004b).

This low expectation of ancient asexuality, though, derives from theory and observations based on macrobes (Normark et al. 2003). Do these expectations apply to all eukaryotes, including microbial lineages? We do not yet know. Little is even known

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about the phylogenetic pattern of asexual microbial eukaryotes. Although many have been postulated to be asexual (Sonneborn 1957), when they have been critically examined, evidence for sex has been found; e.g., Giardia lambli (Birky 2005; Ramesh et al. 2005), and Naegleria lovaniensis (Hurst et al. 1992; Pernin et al. 1992). There are many ways in which microbial eukaryotes could pose challenges to macrobial expectations. For example, many ciliates appear to have globally distributed populations

(Finlay 2002; Foissner et al. 2008), and many also have extremely large population sizes

(Lynch and Conery 2003; Snoke et al. 2006), but see Katz et al. (Foissner et al. 2008;

2006). Such population structures might allow microbial eukaryotes to defy predictions that asexuality might lead to extinction.

If our macrobial theory of low expectations of ancient asexuals does not apply to microbial eukaryotes, then colpodeans may very well be asexual. But if these expectations do apply, then we would expect that colpodeans are covertly sexual since ancient asexuals are not likely.

6.5. Did colpodeans reverse the loss of sex?

If colpodeans are asexual we would have to hypothesize a reverse of the loss of sex. This is because the sexual B. truncatella is nested within putatively asexual clades (Fig. 1:

Dunthorn et al. 2009; Dunthorn et al. 2008; Lynn et al. 1999). Although there is debate on the longevity of ciliate species (Dunthorn and Katz 2008; Nanney 1999), this reversal back to sexuality in B. truncatella could have occurred many millions of years ago.

There is debate about the ability to reverse the loss of a complex character (Bull and Charnov 1985; Collin and Miglietta 2008; Goldberg and Igic 2008; Gould 1970;

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Porter and Crandall 2003; Simpson 1953; Teotónio and Rose 2001). Complex characters can either be lost phenotypically or genotypically; the ability to regain depends on which of these levels was originally involved and the amount of time intervening loss and reversal (Collin and Miglietta 2008). If the genes remain, then regaining a character can just be a process of shuffling around genotypes or turning back on suppressed loci. If the genotype is lost, then it is much harder to regain the lost character.

Can the loss of sex be reversed? Sex may have been lost and regained along lineages throughout the eukaryotic tree of life, though we would not know given the current distribution of sex in extant species (Williams 1975). We do know of two putative cases of regaining sexuality—both at relatively shallow nodes. In multiple populations of the plant Hieracium pilosella (Chapman et al. 2003), reversal from asexuality to sexuality entailed returning to homozygosity (and tetraploidy) for a recessive allele (Bicknell et al.

2000). In this case, although the phenotype of sex was lost, alleles encoding this phenotype remained in the population. In oribatid mites, the case for reversal depends on ancestral character state reconstructions (Domes et al. 2007). However, these ancestral state reconstructions may be fundamentally flawed, leading to a false acceptance of reversal (Goldberg and Igic 2008). The case of regaining sex in oribatid mites is thus ambiguous, and may represent multiple, independent losses of sex. There is no current evidence on whether the genotype was lost in oribatid mites. It should also be noted that had Domes et al. (2007) applied their method to other macrobial taxa they may have increased the number of putative cases of reversing the loss of sex, although these would have been fundamentally flawed as well.

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If sex-related loci are shown to be retained in the putative asexual relatives of B. truncatella a strong case for reversing the loss of sex in colpodeans can be made.

However, the problem of reactivation of silenced loci increases over time as they may be mutated or lost (Collin and Miglietta 2008; Marshall et al. 1994; Normark et al. 2003).

Adding the possible issues surrounding ancient asexuality, though, to the temporal issues of reactivating loci would compound the problem of the colpodeans being asexual. If we assume that both the phenotype and genotype of sex were lost, then our expectations for reversing the loss of sex would be much smaller to none in the colpodeans.

6.6. Conclusion

We will probably not know if colpodeans are sexual until someone actually sees conjugation in a Petri dish and reliably demonstrates nuclear exchange for additional species. The lack of evidence for sex (e.g., no observation of conjugation) is not itself evidence. Given the theories for the maintenance of sex, the combined problems of ancient asexuals and reversing a complex trait, we suggest that many, if not all, colpodeans are covertly sexual, not asexual. Nevertheless, it would be fruitful if it is shown that macrobial theoretical expectations for ancient asexuality apply universally to eukaryotes such ciliates and other microbial eukaryotes. It would also be fruitful to look for loss or retention of sex-related loci in colpodean species.

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Bell, G. 1982. The masterpiece of nature: the evolution and genetics of sexuality. University of California Press, Berkeley.

Bell, G. 1988. Sex and death in protozoa: the history of an obsession. Cambridge University Press, Cambridge.

Bicknell, R. A., N. K. Borst, and A. M. Koltunow. 2000. Monogenic inheritance of apomixis in two Hieracium species with distinct developmental mechanisms. Heredity 84:228-237.

Birky, C. W. 1996. Heterozygosity, heteromorphy, and phylogenetic trees in asexual eukaryotes. Genetics 144:427-437.

Birky, C. W. 2005. Sex: is Giardia doing in the dark? Curr. Biol. 15:R56-R58.

Bull, J. J., and E. L. Charnov. 1985. On irreversible evolution. 39:1149-1155.

Burt, A. 2000. Sex, recombination, and the efficacy of natural selection — was Weisman right? Evolution 54:337-351.

Ceplitis, A. 2003. Coalescence times and the Meselson effect in asexual eukaryotes. Genet. Res. 82:183-190.

Chapman, H., G. J. Houliston, B. Robson, and I. Iline. 2003. A case of reversal: the evolution and maintenance of sexuals from parthenogenetic clones in Hieracium pilosella. Int. J. Plant Sci. 164:719-728.

Collin, R., and M. P. Miglietta. 2008. Reversing opinions on Dollo's Law. Trends Ecol. Evol. 23:602-609.

Dacks, J., and A. J. Roger. 1999. The first sexual lineage and the relevance of facultative sex. J. Mol. Evol. 48:779-783.

Dini, F., and D. Nyberg. 1993. Sex in ciliates. Adv. Microb. Ecol. 13:85-153.

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Domes, K., R. A. Norton, M. Maraun, and S. Scheu. 2007. Reevolution of sexuality breaks Dollo's law. Proc. Natl. Acad. Sci. USA 104:7139-7144.

Dunthorn, M., W. Foissner, and L. A. Katz. 2008. Molecular phylogenetic analysis of class Colpodea (phylum Ciliophora) using broad taxon sampling. Mol. Phylogenet. Evol. 48:316-327.

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Finlay, B. J. 2002. Global dispersal of free-living microbial eukaryote species. Science 296:1061-1063.

Fischer, R. A. 1930. The genetical theory of natural selection. Oxford University Press, Oxford.

Foissner, W. 1985. Klassifikation und phylogenie der Colpodea (Protozoa: Ciliophora). Arch. Protistenk. 129:239-290.

Foissner, W. 1993. Colpodea (Ciliophora). Protozoenfauna 4/1:i-x, 1-798.

Foissner, W., A. Chao, and L. A. Katz. 2008. Diversity and geographic distribution of ciliates (Protista: Ciliophora). Biodivers. Conserv. 17:345-363.

Goldberg, E. E., and B. Igic. 2008. On phylogenetic tests of irreversible evolution. Evolution 62:2727-2741.

Gould, S. J. 1970. Dollo on Dollo's Law: irreversibility and the status of evolutionary laws. J. Hist. Biol. 3:189-212.

Hamilton, W. D. 2001. Narrow roads of gene land, Vol. 2: the evolution of sex. Oxford University Press, Oxford.

Hur, J. H., K. V. Doninck, M. L. Mandino, and M. Meselson. 2008. Degenerate tetraploidy was established before bdelloid rotifer families diverged. Mol. Bio. Evol. 26:375-383.

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Hurst, L. D., W. D. Hamilton, and R. J. Ladle. 1992. Covert sex. Trends Ecol. Evol. 7:144-145.

Judson, O. P., and B. B. Normark. 1996. Ancient asexual scandals. Trends Ecol. Evol. 11:A41-A46.

Katz, L. A., J. G. Bornstein, E. Lasek-Nesselquist, and S. V. Muse. 2004. Dramatic diversity of ciliate histone H4 genes revealed by comparisons of patterns of substitutions and paralog divergences among eukaryotes. Mol. Biol. Evol. 21:555- 562.

Kondrashov, A. S. 1982. Selection against harmful mutations in large sexual and asexual population. Genet. Res. 40:325-332.

Kondrashov, A. S. 1993. Classification of hypotheses on the advantage of amphimixis. J. Hered. 84:372-387.

Lynch, M., R. Bürger, D. Butcher, and W. Gabriel. 1993. Mutational meltdowns in asexual populations. J. Hered. 84:339-344.

Lynch, M., and J. S. Conery. 2003. The origins of genome complexity. Science 302:1401-1404.

Lynn, D. H. 1976. Comparative ultrastructure and systematics of colpodida. Structural conservatism hypothesis and a description of Colpoda steinii Maupas. J. Protozool. 23:302-314.

Lynn, D. H. 2008. The ciliated protozoa: characterization, classification, and guide to the literature, 3rd edition. Springer, Dordrecht.

Mark Welch, D., M. P. Cummings, D. M. Hillis, and M. Meselson. 2004a. Divergent gene copies in the asexual class Bdelloidea (Rotifera) separated before the bdelloid radiation or within bdelloid families. Proc. Natl. Acad. Sci. USA 101:1622-1625.

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Mark Welch, D., J. L. Mark Welch, and M. Meselson. 2008. Evidence for degenerate tetraploidy in bdelloid rotifers. Proc. Natl. Acad. Sci. USA 105:5145-5149.

Mark Welch, D., and M. Meselson. 2000. Evidence for the evolution of bdelloid rotifers without sexual reproduction or genetic exchange. Science 288:1211-1215.

Mark Welch, J. L., D. Mark Welch, and M. Meselson. 2004b. Cytogenetic evidence for asexual evolution of bdelloid rotifers. Proc. Natl. Acad. Sci. USA 101:1618-1621.

Marshall, C. R., E. C. Raff, and R. A. Raff. 1994. Dollo's law and the death and resurrection of gene. Proc. Natl. Acad. Sci. USA 91:12283-12287.

Martín-González, A., J. Wierzchos, J. C. Gutiérrez, J. Alonso, and C. Ascaso. 2008. Morphological stasis of protists in lower Cretaceous amber. Protist 159:251-257.

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Muller, H. J. 1964. The relation of recombination to mutational advance. Mutation Res. 1:2-9.

Nanney, D. L. 1980. Experimental ciliatology: an introduction to genetic and developmental analysis in ciliates. John Wiley & Sons, New York.

Nanney, D. L. 1999. When is a rose? the kinds of Tetrahymena. Pp. 93-118 in R. A. Wilson, ed. Species: new interdisciplinary essays. The MIT Press, Cambridge.

Normark, B. B., O. P. Judson, and N. A. Moran. 2003. Genomic signatures of ancient asexual lineages. Biol. J. Linn. Soc. 79:69-84.

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Ramesh, M. A., S.-B. Malik, and J. M. Longsdon. 2005. A phylogenomic inventory of meiotic genes: evidence for sex in Giardia and an early eukaryotic origin of meiosis. Curr. Biol. 15:185-191.

Schmidt, A. R., H. von Eynatten, and M. Wagreich. 2001. The Mesozoic amber of Schliersee (southern Germany) is Cretaceous in age. Cretaceous Res. 22:423-428.

Schwander, T., and B. J. Crespi. 2009. Twigs on the tree of life? Neutral and selective models for intergrating macroevolutionary patterns and microevolutionary processes in the analysis of asexuality. Molec. Ecol. 18:28-42.

Schönborn, W., H. Dörfelt, W. Foissner, L. Krientiz, and U. Schäfer. 1999. A fossilized microcenosis in Triassic amber. J. Eukaryot. Microbiol. 46:571-584.

Simpson, G. G. 1953. The major features of evolution. Columbia University Press, New York.

Smith, R. J., T. Kamiya, and D. J. Horne. 2006. Living males of the 'ancient asexual' Darinulidae (Ostracoda: Crustacea). Proc. R. Soc. Lond. B 273:1569-1578.

Snoke, M. S., T. U. Berendonk, D. Barth, and M. Lynch. 2006. Large global effective population sizes in Paramecium. Mol. Biol. Evol. 23:2474-2479.

Teotónio, H., and M. R. Rose. 2001. Perspective: reverse evolution. Evolution 55:653- 660.

West, S. A., C. M. Lively, and A. F. Read. 1999. A pluralist approach to sex and recombination. J. Evol. Biol. 12:1003-1012.

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Figure 6.1. Phylogeny and distribution of sex within the Colpodea. Major clades are labeled. Only Bursaria truncatella is known to have sex. The ancestral state (?) for colpodeans is debated: either sexual or asexual. Modified from Dunthorn et al. (2009).

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CHAPTER 7

PHYLOCODE DEFINITIONS FOR FOUR CILIATE CLADES

Micah Dunthorna and Denis H. Lynnb

aGraduate Program in Organismic and Evolutionary Biology, University of

Massachusetts, Amherst, Massachusetts, 01003, USA bDepartment of Integrative Biology, University of Guelph, Guelph, Ontario, N1G 2W1,

Canada

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7.1. Abstract

Ciliates clades have traditionally been named using the International Code of

Zoological Nomenclature, a rank-based system that governs names at or below the family rank. Here we argue that the Internal Code of Phylogenetic Nomenclature should be used to name ciliate. We apply this code to four ciliate clades above the rank of family:

Ciliophora, Postciliodesmatophra, Intramacronucleata, and Colpodea.

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7.2. Introduction

Names for ciliate taxa are currently governed by the International Code of

Zoological Nomenclature (International Commission on Zoological Nomenclature 1999); hereafter referred to as the ICZN. Here I will briefly argue that ciliate taxa should be named using the PhyloCode, which is governed by the International Code of

Phylogenetic Nomenclature (Cantino and de Queiroz 2007); hereafter referred to as the

ICPN. As an example, four well-known ciliate names are converted using the ICPN.

Ciliates are a clade of microbial eukaryotes with dimorphic nuclei and cilia in at least one life-cycle stage (Lynn 2008). The diploid ‘germline’ micronucleus is, for the most part, transcriptionally inactive and is exchanged between ciliates during sex. The usually polyploid ‘somatic’ macronucleus is transcriptionally active (Raikov 1996).

Ciliate morphospecies are described by the patterns formed by the kineties (arrays) of kinetosomes (cilia basal bodies) and associated fibers in the somatic and oral membranes.

Analyses of these rich characters have led to several taxonomic schemes at a variety of levels in the ciliate tree of life (Bütschli 1887-1889; Kahl 1926, 1930-1935; Kahl 1931;

Puytorac et al. 1974; Corliss 1979; Small and Lynn 1981; Foissner 1985; Matthes 1988;

Foissner 1993; Puytorac 1994; e.g., Berger 1999; Lynn and Small 2002; Berger 2006;

Lynn 2008).

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7.3. The PhyloCode

The ICPN is a nomenclatural system that explicitly uses the theoretical approach of phylogenetic systematics. In essence, this system uses hypotheses of relationships, or distributions of apomorphies to name clades (monophyletic groups). To validly name a taxon using the ICPN there must be a description of clade or species using internal and, sometimes, external, specifiers that are either species or characters (Cantino and de

Queiroz 2007). Ideally there should be at least one reference phylogeny that includes the specifiers that accompanies the description (Cantino and de Queiroz 2007). ICPN names can be converted from the ICZN, or other rank-based codes, or new names can be coined.

Three main types of definitions are used in the ICPN: node-based, branch-based, and apomorphy-based(Cantino and de Queiroz 2007). The exact wording for each definition can take many forms, and node-based definitions can be branch- and apomorphy-modified. Which type of definition to use depends on a number of issues; e.g., the presence of absence of basal fossil lineages, the amount of support for the monophyly of the clade, support for the relationships within the clade, or support for sister groups (Bryant 1994; Lee 1996; Sereno 1999, 2005; Cantino and de Queiroz 2007;

Cantino et al. 2007).

A phylogenetically-based nomenclature system like the ICPN, as in any system, has its strengths (de Queiroz and Gauthier 1990; de Queiroz and Gauthier 1992, 1994;

Cantino et al. 1997; de Queiroz 1997; Lee 2001; Bryant and Cantino 2002; Pleijel and

Rouse 2003; Sereno 2005; de Queiroz 2006; Laurin et al. 2006; de Queiroz 2007) and weaknesses (Dominguez and Wheeler 1997; Moore 1998; Benton 2000; Nixon and

Carpenter 2000; Dyke 2002; Carpenter 2003; Keller et al. 2003; Kojima 2003; Moore

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2003; Nixon et al. 2003; Schuh 2003; Barkley et al. 2004; Wenzel et al. 2004; Polaszek and Wilson 2005). Only a few of these strengths will be briefly discussed below to show why the ICPN should be used to name ciliate clades.

7.4. Why the PhyloCode should be applied to ciliates

The ICPN has been used in a number of terminal, macrobial eukaryotic taxa such as animals, plants, and fungi (Hibbett and Donoghue 1998; Donoghue et al. 2001;

Stefanovic et al. 2003; Joyce et al. 2004; Modesto and Anderson 2004; Sangster 2005;

Taylor and Naish 2005; Cantino et al. 2007). Recently, Adl et al. (2007) argued that microbial eukaryote species should be named using the ICPN or a similar nomenclatural system, but they did not call for its application in naming deeper microbial eukaryotic clades. Here we argue that the ICPN should be applied to deeper microbial eukaryotic nodes, using ciliates as an example.

The ICPN applies at all phylogenetic depths. Deep nodes in the ciliate tree of life are not covered by the ICZN because they are above the family rank. There are attempts to make the ICZN and other rank-based codes apply at higher ranks (Hemming 1953;

Corliss 1972; Ghiselin 1977; Brothers 1983; Dundee 1989; Dubois 2005, 2006), but these suggestions have yet to be codified. Those using the ICZN, or any rank-based system, above the family rank are doing so with the consensus of the authors and users of the classification, not with authority of these codes (Corliss 1983). On the other hand, the

ICPN governs the name of any taxon at any depth in the tree of life.

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The ICPN allows for only names that designate taxa that are hypothesized to be monophyletic. The ICZN does not say anything about what kind of groups

(monophyletic, paraphyletic, or polyphyletic) can or cannot be named. There is opposition to strictly monophyletic taxonomies (Sosef 1997; Mayr 1998; Thorne 2000;

Mayr and Bock 2002; Wu et al. 2002; Brummitt 2003; Nordal and Stedje 2005; Heywood et al. 2007; Hörandl 2007), but there still is no way to easily interpret paraphyletic or polyphyletic taxa. The majority of macrobial classifications only recognize monyphyletic taxa (e.g., APG II 2003; Faivovich et al. 2005), athough paraphyletic taxa are still recognized researchers for microbial eukaryotes in general (e.g., Cavalier-Smith 1999;

Cavalier-Smith 2004, 2007) and ciliate in specific (e.g., Berger 1999, 2006).

Monophyletic taxa sensu Hennig (1966) or Farris (1974) are unambiguous, reflect sister- group relationships that are objectively stated and interpreted. Although the ICPN does not prevent the recognition of paraphyletic and polyphyletic taxa (de Queiroz 2006), it does restrict the application of names to only groups that are hypothesized to be monophyletic.

In the ICPN named ranks will not necessarily occur. In the ICZN, a family, genus, or species name is only valid if it is given a rank, and the name may change if its rank is changed. Ranked names are neither mandated nor prohibited by the ICPN (de Queiroz

2006; Cantino and de Queiroz 2007). Even if an ICPN name has a ranked ending, a change in name will not occur if hypotheses of relationships are revised later.

The ICPN allows for explicit statements of when a name should be rejected and no longer used through explicit statement about which taxa can be included and excluded.

Although taxonomies are often stated as hypotheses, there is usually no way to know

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when the name should be rejected and no longer used as in the ICZN unless there is a nomenclatural act later on. The ICPN provides a system in which a taxon name can be explicated stated when it should be used and when it should abandoned in the light of new data through the use of specifiers and qualifying clauses (Schander and Thollesson

1995; Cantino et al. 1997; Sereno 1999; Bryant and Cantino 2002; Joyce et al. 2004;

Sereno 2005; Bertrand and Härlin 2006).

7.5. Application of the PhyloCode to four ciliate clades

Below are four ciliate clades to which we apply the ICPN. Since they are yet to be published in the International Code of Phylogenetic Nomenclature’s Companion Volume, which will be the official start for all valid PhyloCode names and definitions, because theses names have not been reviewed by outside taxonomists, and because these definitions are not published in an easily attainable journal, the definitions below are neither effectively nor validly published here. The entire ciliate clade (currently ranked at the phylum level in the ICZN) and the Postciliodesmatophora and Intramacronucleata

(the two currently subphyla) and the Colpodea (ranked as a class) are converted here. In all definitions ranks are ignored. Taxa all italicized following standard formatting of the

ICPN.

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7.5.1. Ciliophora

F. Doflein 1901 [M. Dunthorn & D. H. Lynn 2009], converted clade name

Definition: The least inclusive clade containing Tetrahymena thermophila Nanney and

McCoy 1976, Blepharisma americanum Suzuki 1954, and Loxodes striatus (Engelmann

1862). This is a node-based definition in which all of the specifiers are extant; it is intended to apply to a crown clade. Abbreviated definition: < Tetrahymena thermophila

Nanney and McCoy 1976 & Blepharisma americanum Suzuki 1954 & Loxodes striatus

(Engelmann 1862).

Etymology: Derived from the Latin cilium (eyelash) and Greek phoreus (bearer), in reference to the cilia on the cell bodies of all members of this clade (in at least one stage of the life cycle).

Reference Phylogeny: The primary reference phylogeny is Hammerschmidt et al. (1996:

Fig. 2). See also Baldauf et al. (2000: Fig. 1) and Yoon et al. (2008: Fig. 2).

Composition: Almost all known ciliates are extant; those few fossils found can be placed within previously recognized taxa (Lynn 2008). The clade Ciliophora contains

Postciliodesmatophora and Intramacronucleata as defined in this volume, which in turn include all of the clades listed by Lynn (2008).

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Diagnostic Apomorphies: Ciliates have three apomorphic characters: 1) dimorphic nuclei, with a “germline” micronucleus and a “somatic” macronucleus that are not homologous with those found in Foraminifera Lee 1990; 2) cilia, in at least one life-cycle stage, that are derived from a kinetosome (= eukaryotic basal body) that is associated with three fibers (a kinetodesmal fiber, a postciliary microtubular ribbon, and a transverse microtubular ribbon); and 3) sex in the form of conjugation, where there is typically mutual exchange of haploid meiotic products of the micronucleus (Raikov 1996; Lynn

2008).

Synonyms: Ciliata M. Perty 1852 [approximate], Infusoria Bütschli 1887 [approximate]; see review by Lynn (2008).

Comments: Ciliates have long been recognized as a group because of their distinctive morphology, and molecular data strongly support this clade (Hammerschmidt et al. 1996;

Lynn 2008). Since the beginning of the 20th century, the name Ciliophora has been the most widely used (e.g., Corliss 1979; Puytorac 1994; Jankowski 2007; Lynn 2008).

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7.5.2. Postciliodesmatophora

Z. P. Gerassimova & L. N. Seravin 1976 [M. Dunthorn & D. H. Lynn 2009], converted clade name

Definition: The least inclusive clade containing Blepharisma americanum

Suzuki 1954 and Loxodes striatus (Engelmann 1862). This is a node-based definition in which both specifiers are extant; it is intended to apply to a crown clade. Abbreviated definition: < Blepharisma americanum Suzuki 1954 & Loxodes striatus (Engelmann

1862).

Etymology: Derived from the Latin post (after, behind) and cilium (eyelash) and the

Greek desmos (bond or chain) and phoreus (bearer), in reference to the postciliodesmata borne by members of this clade (see Diagnostic Apomorphies).

Reference Phylogeny: The primary reference phylogeny is Hammerschmidt et al. (1996:

Fig. 2). See also Hirt et al. (1995: Fig. 2).

Composition: Contains the Karyorelictea and the Heterotrichea; see Lynn (Lynn 2008) for a list of the contents of these clades.

Diagnostic Apomorphies: The Postciliodesmatophora have postciliodesmata, formed from postciliary microtubule ribbons overlapping along the right side of a kinety

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(integrated somatic file of kinetids). Macronuclei either do not divide (Karyorelictea) or divide using extra-macronuclear microtubules (Heterotrichea).

Synonyms: None.

Comments: Gerassimova and Seravin (1976) recognized this clade based on the shared postciliodesmata. Subsequent phylogenetic analyses of small subunit ribosomal RNA genes have supported its monophyly (Hirt et al. 1995; Hammerschmidt et al. 1996; Lynn

2008). The name is defined here in a manner consistent with the intent of the original conception of the group by Gerassimova and Seravin (1976) as containing only

Karyorelictea and the Heterotrichea. If a lineage is found that has postciliodesmata but falls below the divergence between Karyorelictea and Heterotrichea, this larger clade will have to have another name.

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7.5.3. Intramacronucleata D. H. Lynn 1996 [D. H. Lynn & M. Dunthorn 2009], converted clade name

Definition: The most inclusive crown clade exhibiting intramacronuclear microtubules

(as described under Diagnostic Apomorphies) synapomorphic with those in Tetrahymena thermophila Nanney and McCoy 1976. This is an apomorphy-modified node-based definition in which the specifier is extant; it is intended to apply to a crown clade.

Abbreviated definition: >∇ exhibiting intramacronuclear microtubules (Tetrahymena thermophila Nanney and McCoy 1976).

Etymology: Derived from the Latin intra (within), Greek makros (large), and Latin nucleus (kernel), in reference to the presence of intramacronuclear microtubules during cell division (see Diagnostic Apomorphies).

Reference Phylogeny: The primary reference phylogeny is Hammerschmidt et al.

Hammerschmidt et al. (1996: Fig. 2). See also Hirt et al. (1995: Fig. 2) and Yoon et al.

(2008: Fig. 2).

Composition: The majority of clades within the Ciliophora, as defined in this volume, are within the Intramacronucleata. Lynn (2008) lists the following taxa as included in the

Intramacronucleata: Armophorea, Colpodea, Litostomatea, Nassophorea,

Oligohymenophorea, Phyllopharyngea, Plagiopylea, Prostomatea, and Spirotrichea.

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Diagnostic Apomorphies: The most distinctive diagnostic apomorphy is the presence of microtubules within the macronuclear envelope during nuclear division. In other

Ciliophora (i.e. Postciliodesmotophora), microtubules are either extra-macronuclear, or macronuclei do not divide.

Synonyms: None

Comments: Lynn (1996) established this taxon, which contains most known ciliates, based on small subunit ribosomal RNA gene phylogenies as well as the presence of intranuclear microtubules in dividing macronuclei. Monophyly of this clade is strongly supported, but internal relationships are unresolved (Lynn 2008).

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7.5.4. Colpodea E. B. Small & D. H. Lynn 1981 [M. Dunthorn & D. H. Lynn 2009], converted clade name

Definition: The most inclusive clade exhibiting a LKm fiber (as described below under

Diagnostic Apomorphies) synapomorphic with that in Colpoda cucullus O. F. Müller

(1773) K. C. Gmelin 1790. This is an apomorphy-modified node-based definition in which the specifier is extant; it is intended to apply to a crown clade. Abbreviated definition: >∇ exhibiting a LKm fiber (Colpoda cucullus O. F. Müller (1773) K. C.

Gmelin 1790).

Etymology: Derived from the Greek kolpus (womb).

Reference Phylogeny: The primary reference phylogeny is Dunthorn et al. (2008: Fig. 2).

See also Dunthorn et al. (2009), and Lynn et al. (1999).

Composition: All taxa listed in Foissner (1993) and Lynn (2008).

Diagnostic Apomorphies: The most distinctive diagnostic apomorphy of the Colpodea is the LKm fiber (=transversodesmal fiber) composed of overlapping transverse ribbons of microtubules extending from the posterior kinetosome (= eukaryotic basal body) of the somatic dikinetids (Foissner 1993; Lynn 2008). The Colpodea also have: somatic stomatogenesis, where parental oral structures are partially to completely reorganized during cell division; and a reticulate silverline system.

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Synonyms: None.

Comments: Using the structural conservatism hypothesis (Lynn 1976, 1981), Small and

Lynn (1981) brought once disparate taxa together into the Colpodea. The Colpodea was expanded by Foissner (1985), who later monographed the group (Foissner 1993). Bardele

(1989) rejects the monophyly of the taxon based on the presence or absence of ciliary plaques, but there is no support for this claim (Lynn et al. 1999; Dunthorn et al. 2008).

Monophyly of the Colpodea is currently neither supported nor rejected by small subunit ribosomal DNA sequences (Dunthorn et al. 2008).

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