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Tardigrade evolution and ecology

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Tardigrade evolution and ecology
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Nichols, Phillip Brent
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Ecdysozoa
Meiofauna
Phylogeny
18s rrna
Morphology
Dissertations, Academic -- Biology -- Doctoral -- USF
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ABSTRACT: A character data set suitable for cladistic analysis of tardigrades at the family level was developed. The data matrix consisted of 50 morphological characters from 15 families of tardigrades and was analyzed by maximum parsimony. Kinorhynchs, loriciferans and gastrotrichs were used as outgroups. The results agree with the currently accepted hypothesis that Eutardigrada and Heterotardigrada are distinct monophyletic groups. Among the eutardigrades, Eoyhypsibiidae was found to be a sister group to Macrobiotidae + Hypsibiidae, while Milnesiidae was the basal eutardigrade family. The basal heterotardigrade family was found to be Oreellidae. Echiniscoideans grouped with some traditional Arthrotardigrada (Renaudarctidae, Coronarctidae + Batillipedidae) suggesting that the arthrotardigrades are not monophyletic. An 18S rRNA phylogenetic hypothesis was developed and supports the monophyly of Heterotardigrada and of Parachela versus Apochela within the Eutardigrada. Mapping of habitat preference suggest that terrestrial tardigrades are the ancestral state. Molecular analysis of a sediment sample with an unusually large population of tardigrades had a higher diversity when compared to manual sorting and counting.
Thesis:
Dissertation (Ph.D.)--University of South Florida, 2005.
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Includes bibliographical references.
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by Phillip Brent Nichols.
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Includes vita.

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Tardigrade Evolution And Ecology by Phillip Brent Nichols A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy Department of Biology College of Arts and Sciences University of South Florida Major Professor: James R. Garey, Ph.D. Richard P. Wunderlin, Ph.D. Florence M. Thomas, Ph.D. Frank A. Romano, III, Ph.D. Date of Approval: July 25, 2005 Keywords: ecdysozoa, meiofauna, phylogeny, 18s rrna, morphology Copyright 2005 P. Brent Nichols

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Dedication For MaMa…

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Acknowledgements There are so many people th at I wish to thank and acknowledge for their love, support, and help over the past few years. Firs t of all to the most important person in my life, my wife Tanya, you are my light and my inspiration, thank you for your unwavering love and support. My major advisor, Dr. James R. Garey, has been a wonderful mentor who has helped me see the “Big Picture” on mo re than one occasion. I will always value both the professional and personal friendship th at we have. I would like to thank the members of my Graduate Supervisory Co mmittee, Dr. Richard P. Wunderlin, Dr. Florence M. Thomas, and Dr. Frank A. Romano, III for their participation in my graduate education and training and a special thank you to Dr. Rick Oches for acting as my defense chair. The members of the Garey Lab; Terry Campbell, Hattie Wetherington, Kim Fearne, John Slomba, Mike Robeson, H eather Hamilton, and Stefie Depovic; have been involved with many aspects of this di ssertation from collecting field samples to problem solving data collection. Other colla borators that have been influential in the completion of my graduate training include : Dr. Diane Nelson, East Tennessee State University; Dr. Ruth Dewel, Appalachian State University; Dr. Roberto Guidetti, University of Modena, Modena, Italy; Dr Reinhardt Kristensen, University of Copenhagen, Denmark; Dr. Sandra McInnes, Cambridge University; Mr. Nigel Marley, University of Plymouth; Mr. Ken Hayes, Univ ersity of Hawaii. I would like to extend my appreciation to the University of South Florida Gradua te School and Biology Department not only for financial support thr oughout my training but for all the “little things” that often go unnoticed. Lastly, I would like to th ank my parents for providing me with the opportunities to pur sue my own interests.

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i Table of Contents List of Tables iii List of Figures iv Abstract vi Chapter One – Introduction 1 Systematic Research 5 Dispersal 9 Ecological Importance 10 Significance 11 Objectives 13 Chapter Two – Specimen Acquisition and Processing 14 Chapter Three – Morphological Analysis of Tardigrade Phylogeny 15 Background 15 Material and Methods 16 Characters 16 Data Analysis 17 Results & Discussion 17 Morphological Phylogeny 17 Habitat Mapping 19 Chapter Four – Molecular Analysis of Tardigrade Phylogeny 20 Background 20 Material and Methods 22 Gene Selection 22 DNA Extraction 23 Polymerase Chain Reaction 24 Cloning 24 Sequencing 25 Alignments and Sequence Analysis 25 Results & Discussion 26 Molecular Phylogeny 26 Habitat Mapping 29

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ii Chapter Five –Meiofaunal abundance, diversity and similarity in a uniquely rich tardigrade community. 31 Background 31 Material and Methods 34 Sample Collection 34 DNA Sediment Extraction 34 Polymerase Chain Reaction 35 Cloning 36 Sequencing 36 Alignments and Sequence Analysis 37 Sediment Sorting and Specimen Id entification 37 Data Analysis 37 Results & Discussion 38 Species Diversity 48 Community Similarity 48 Chapter Six– Summary 51 Morphological Analysis of Tardigrade Phylogeny 51 Molecular Analysis of Tardigrade Phylogeny 51 Meiofaunal Abundance, Diversity and Similarity in a Uniquely Rich Tardigrade Community 52 References 53 Bibliography 62 Appendices 86 Appendix A. Morphological Data Matrix 88 Appendix B. List of Morphologica l Characters 89 About the Author End Page

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iii List of Tables Table 1. Modes and causes of crytobiotisis in tardigrades. 4 Table 2*. Higher level Taxonomy of the Ta rdigrada. 9 Table 3. List of species, sequence length, orig in, and sequence reference. 26 Table 4. Sequence groups and the nu mber of sequences per group. These data were used to calculat e Shannon diversity, Simpson’s Diversity, Jaccard’s Similarity, and Stander’s Similarity. 39 Table 5. Species groups and frequency of individuals per group. These data were used to calculate Sh annon diversity, Simpson’s Diversity, Jaccard’s Similarity, and Stander’s Similarity. 47 Table 6. Shannon (H’) and Simpson’s (C ) diversity indices for molecular OTU data and species groups. 48 Table 7. Jaccard’s and Stander’s commun ity similarity indices for molecular OTU community compared to the manual count community. 49

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iv List of Figures Figure 1. Graphical representation of a t ypical tardigrade Redrawn from Rammazzotti and Maucci (1983). 1 Figure 2. Typical tard igrade claws. A. Macrobiotus sp.; B. Milnesium sp.; C. Echiniscus sp.; D. Batillipes sp. Redrawn from Rammazzotti and Maucci (1983). 5 Figure 3. Representative head struct ures of eutardigrades (left) and heterotardigrades (right). A. Cephalic papillae; B. Peribuccal papillae; C. Eye spots; D. Internal cirrus; E. Cephalic papilla; F. External cirrus; G. Late ral Cirrus A; and H. Clava. Redrawn from Rammazzotti and Maucci (1983). 6 Figure 4. Typical buccal apparatus found in tardigrades. A. Stylet; B. Buccal tube; C. Stylet support; D. M acroplacoid; E. Microplacoid; F. Pharyngeal bulb Redrawn from Rammazzotti a nd Maucci (1983). 7 Figure 5. Maximum parsimony phylogeny of tardigrade families with mapped habitat preferences. The analysis supports the monophyletic origin of the class Eutardigrada and or ders Parachela and Apochela along with the class Heterotardigrada; however, the orders within Heterotardigrada (Arthrotardigr ada, Echiniscoidea) may not be monophyletic. Key to Characters: 1. Eutardigrada: No cephalic appendages; lack of dorsal plates ; differentiated placoids; double claws with secondary and primar y branches. 2. Heterotardigrada: Cephalic appendages, digitate le gs with or without claws. 3. Parachela: No cephalic papill ae; double claws per leg divided into a secondary and primary bran ch. 4. Apochela: Six cephalic papillae; two sets of claws per leg, with unbranched primary claw separated from secondary claw; 4 6 peribuccal lamellae. Habitats: Terrestrial (T); Marine (M); or Freshwater (F). Maximum parsimony bootstrap values are shown. 18

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v Figure 6. Topology of an 18S rRNA ba sed tree mapped with diagnostic morphological characters (Gar ey et al. 1999). The molecular tree is completely congruent w ith current morphological hypotheses of tardigrade phylogeny. Bootst rap and confidence probability values are for NJ and MP trees ar e shown at each node. Key to Characters: 1. Arthropoda + Tardigrada: supraesophageal or preoral position of the frontal appendages and their neuromeres (DEWEL & DEWEL 1997). 2. Arthropoda: body with articu lated exoskeleton; protocerebrum with compound eyes (NIELSEN 1995) 3. Tardigrada: connective between protocerebrum and ganglion of first pair of legs DEWEL & DEWEL 1997). 4. Heterotardigrada: Cephalic appendages, legs with digits and/or claws (BARNES & HARRISON 1993). 5. Eutardigrada: Lack of cephalic appendages; legs with claw s but not digits (B ARNES and HARRISON 1993). 6. Apochela: With cephali c papillae and with double claws with well-separated primary and sec ondary branches (SCHUSTER et al. 1980). 7. Parachela: Without cephalic papillae and with double claws in which primary and secondary branch es are joined (SCHUSTER et al. 1980). 8. Macrobiotus: Claw branches with sequence: secondary, primary, primary, secondary; buccal tube wi th ventral lamina and 10 peribuccal lamellae (SCHUSTER et al. 1980 ). 9. Thulinia + Hypsibius: Claw branches with sequence: sec ondary, primary, secondary, primary; buccal tube without ventral lamina (SCHUSTER et al. 1980; BERTOLANI 1982). 10. Thulinia: twelve peri buccal lamellae (BERTOLANI 1982). 11. Hypsibius: peribuccal lamellae abse nt (SCHUSTER et al. 1980). 22 Figure 7. This study used the 18S rRNA gene using primers to amplify the portions of each gene as shown. See text for details 23 Figure 8. Tardigrade phylogeny based on l8S rRNA gene sequences. Similar trees were recovered for maximum parsimony and maximum likelihood analyses. Branch lengths are drawn to scale (substitutions/site). The numb ers above each fork are bootstrap values for the NJ tree. 27 Figure 9. Tardigrade molecular topology from Fig. 8 with mapped habitat preference. Habitats: Terrestrial (T ); Marine (M ); or Freshwater (F ) 30 Figure 10. Location of sedime nt collection site on Dauphin Island, AL. 34 Figure 11. Rarefaction curve calculated for molecular OTUs and species groups. 39

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vi Figure 12. Neighbor-joining phylogenetic tr ee based on the number of differences between sequences and complete deletion of gaps of all 126 sequences. Groups notated to the right of each group consist of sequences with no no more than 5 differences between them. The scale bar on the last page of the tree indicates the number of differences per length of the bar. 41 Figure 13. Neighbor-joining phylogenetic tree of the unique OTU sequences and a reference data set. The phyloge netic tree was created based on the Kimura 2-parameter distance method and complete deletion of gaps. 45

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vii Tardigrade Evolution and Ecology Phillip Brent Nichols ABSTRACT A character data set suitable for cladisti c analysis of tardigrades at the family level was developed. The data matrix cons isted of 50 morphologi cal characters from 15 families of tardigrades and was analyzed by maximum parsimony. Kinorhynchs, loriciferans and gastrotrichs were used as out groups. The results agree with the currently accepted hypothesis that Eutardigrada and He terotardigrada are distinct monophyletic groups. Among the eutardigrades, Eoyhypsibiidae was found to be a sister group to Macrobiotidae + Hypsibiidae, while Milnesiidae was the basal eutardigrade family. The basal heterotardigrade family was found to be Oreellidae. Echiniscoideans grouped with some traditional Arthrotardigrada (Renauda rctidae, Coronarctidae + Batillipedidae) suggesting that the arthrota rdigrades are not monophyletic. An 18S rRNA phylogenetic hypothesis was developed and supports the monophyly of Heterota rdigrada and of Parachela versus Apochela within the Eu tardigrada. Mapping of habitat preference suggest that terrestria l tardigrades are the ancestral st ate. Molecular analysis of a sediment sample with an unusually large popu lation of tardigrades had a higher diversity when compared to manual sorting and counting.

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1 Chapter One Introduction Tardigrades were first described by Goeze in 1773 and since then, his "Kleiner Wasser Brs" have been commonly referred to as “water-bears” because of their bear-like appearance. Spallanzani (1776) termed hi s similar organism "Il Tardigrado" (slow stepper) recognizing the unique lumbering gate exhibited by tardigrades. Tardigrades are found throughout the world in a variety of freshw ater, marine, and terre strial habitats and are considered cosmopolitan in their dist ribution. These bilaterally symmetric, hydrophilous micrometazoans are ventrally flat tened and dorsally convex. A typical adult tardigrade (Figure 1) is 250500 micrometers in length and displays limited metamerism with five indistinct segments. A cephalic segment that is bluntly rounded contains a mouth and may have eyespots and sensory cirri. Four body segments are present, each has a pair of ventrolatera l legs terminating in claws or suction discs (Ramazzotti and Maucci, 1983); generally the first three pair s are used for locomotion, the fourth for substrate attachment. Tardigrades have an out er cuticle that may be opaque, white, or Figure 1. Graphical representation of a typical tardigrade. Redrawn from Ramazzotti and Maucci (1983).

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2 such colors as brown, green, pink, red, or ange, or yellow, covering them (Dewel et al. 1993). This color results from pigments in the cuticle, dissolved materials in the body fluids, or from the contents of the digestive tract. Terrestrial tardigrades mainly feed on algae, cryptogams (mosses, lichens, and liverworts) or animals (rotifers, nematodes, and other small invertebrates). Marine tardigrades are believed to feed primarily on bacteria (R.M. Kristensen, personal communication). Feeding is usually accomplishe d by piercing the cells with a pair of stylets and “sucking” out their contents, but in some cases whole orga nisms are ingested. The typical digestive system is comprised of a foregut, midgut, and hindgut. The foregut includes a mouth, buccal tube, salivary gla nds, stylets, a sucking pharynx (with or without placoids), and an esophagus. The styl ets are extended to pierce the cells, and the pharynx pumps the cytoplasmic fluids into the esophagus. The salivary glands are believed to secrete new stylets during ecdysis (Walz, 1982; Ramazzotti and Maucci, 1983). Food is digested in the midgut and the excretory glands (M alpighian tubules) empty into the junction of the midgut and hindgu t. The hindgut can either have a true cloaca (Eutardigrada) or an anus with a separate, preanal gonopore (Heterotardigrada). Doyere proposed that al l tardigrades were herma phroditic based on what he identified as structures that were two testes and a semi nal receptacle (Bertolani, 1979, Nelson, 1982). This belief remained until two st ructures were identified as malpighian tubules and tardigrades were then consid ered to be gonochoristic (Bertolani, 1992; Nelson 1982). However, Bertolani (1979) a nd Bertolani and Mani cardi (1986) reported that hermaphroditism exists in some tardigrades.

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3 The major modes of reproduction in tardigrades are amphimixis and parthenogenesis (Ramazzotti and Maucci, 1983; Nelson, 1982; Bertolani, 1992; Kinchin, 1994; Bertolani and Rebecchi, 1998). The fema les lay eggs outside the body or within the exuvium as they molt. The males then release spermatozoa in to the exuvium where external fertilization occurs or into the gonopore or cloaca where they will travel up the oviduct for internal fertilization (Pollock, 1975). Sexual reproduction in g onochoristic tardigra des (amphimixis) can take place between a female and a single male or seve ral males with the males clinging to the anterior part of the female with their front legs (Kinchin, 1994). There have been limited investigations into the mating behavior of tardigrades. In stead, mating habits have been inferred from anatomical stud ies (Bertolani, 1992) Males and females are quite similar but can be distinguished from one anot her by comparing the gonad and the gonoducts. Females have one ovoduct whereas males have 2 va s deferens. Fertilization for terrestrial species usually occurs inside the female’s body while in mari ne species fertilization is external (Bertolani, 1990). Parthenogenesis is the development of an egg where there has been no paternal contribution of genes (Futuyma, 1986). Th is is the common mode of reproduction in unisexual tardigrades found in non-marine habita ts (leaf litter, mosses, and freshwater) (Nelson, 1982). Unisexual females are wide sp read in tardigrades but recombination may be limited as they will have fewer possibili ties for procreation. Parthenogenesis, while limiting genetic variability, may in fact faci litate proliferation in unisexual females (Bertolani, 1987). Polyploidy, possessing more than one entire chromosomal compliment has often been associated w ith parthenogenesis and several species are known to have polyploid populations (Bertola ni, 1982; Nelson, 1982). Also, cytotypes,

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4 or populations with the same morphologica l characteristics but differing degrees of ploidy (diploidy, triploidy, and tetraploidy), have been observed from samples collected relatively close to one another (Rebecchi and Bertolani, 1988). The nervous system is composed of a br ain with two dorsolate ral lobes connected by two circumpharyngeal commissures to a subpharyngeal ganglion, a pair of longitudinal nerve cords, and four ventral ganglia that ar e united by those nerve strands (Dewel and Dewel, 1996). The most fascinating feature of some tard igrades is their capacity to enter into a state of suspended animation (cryptobiosis) The water content of the body is reduced from 85% to just 3% and the body becomes barrelshaped forming a tun. In this state, growth, reproduction, and metabolis m are reduced or cease temporarily and resistance to environmental extremes is evident. This re sistance allows the tardigrade to survive through cold and dry spells, ionizing radi ation, heat, and pollution. Crowe (1975) identified five different type s of cryptobiosis (Table 1). Table 1. Modes and causes of crytobiotisis in tardigrades MODE Description Encystment Most common among aquatic and soil tardigrades, they encase themselves by molting and remaining inside the old cuticle. Anoxybiosis Induced by low oxygen environments Cryobiosis Induced by low temperatures Osmobiosis Induced by elevated osmotic stress Anhydrobiosis Induced by the removal of water from the tardigrade and its environment through evaporation.

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5 Systematic Research In 1928 and 1929 Marcus compiled the first reviews of tardigrade morphology, physiology, embryology, and phylogenetic relations hips; in 1936, he developed the first basis for classification of the group (Kinch in, 1994). Ramazzotti's first monograph in 1962, a revision in 1972, and a subsequent revision by Ramazzotti and Maucci in 1983 included descriptions of 514 species in three classes, Heterotardigra da, Eutardigrada, and Mesotardigrada. Mesotardigrada wa s based on a single description of Thermozodium esakii from a hot spring near Nagasaki Japan. Ty pe specimens were not preserved and the hot spring where they were found was destroyed in an earthquake (Ramazzotti and Maucci, 1983). Since the publication of the 1983 monograph, the number of described species has increased to over 900. Modern taxonomy is based on these and a number of other European studies. The history of tardigrade studies is well doc umented by Ramazotti and Maucci (1983) and Kinchin (1994). Increased interest in tard igrade biology and systematics over the last 25 years is evidenced by numerous publicat ions and 9 international symposia. Tardigrade systematics has historically been based on a number of morphological characteristics. The current taxonomy of clades within Tardigrada is based on Figure 2. Typical tardigrade claws. A. Macr obiotus sp.; B. Milnesium sp.; C. Echiniscus sp.; D. Batillipes sp. Redrawn from Ramazzotti and Maucci (1983).

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6 morphological characters that include: claw size, shape, orga nization, and number (Figure 2); organization of the bucco-pharyngeal apparatus; lengt h of stylets; size, shape, and number of placoids; cuticular patterns a nd ornamentation; and morphology of eggs. The presence or absence of the Lateral Cirrus A is used to separate the two major classes, Heterotardigrada and Eutardigrada (Figure 3). The Heterotardigrada are characterized by 4 si ngle claws per leg that may have spurs at the base of the exterior or interior claws. They often have sensory papilla or a spine and/or a dentate collar on th e 4th pair of legs. The Hetero tardigrada are “armored” and have a thick cuticle, dividing into plates, w ith species specific pore patterns. Two double claws per leg, which may or may not be si milar in size and shape, characterize the Eutardigrada. Each double claw consists of a primary and secondary branch and a base. The sequence of branching and other claw ch aracteristics are taxonomically significant. Figure 3. Representative head structures of eu tardigrades (left) and heterotardigrades (right). A. Cephalic papillae; B. Peribuccal pap illae; C. Eye spots; D. Internal cirrus; E. Cephalic papilla; F. External cirrus; G. Lateral Cirrus A; and H. Clava. Redrawn from Ramazzotti and Maucci (1983).

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7 The eutardigrades have a thin cuticle that is not divided into plates, but may be ornamented with spines, reticu lations, granules, or pores. The tardigrade buccal apparatus consists of a pair of piercing stylets, a buccal tube, and a sucking pharynx. Peribuccal papillae, papulae or lobes (Schuster et al ., 1980) may be present around the mouth in eutardigra des. The opening of the mouth is followed by the buccal tube which is supported by a ventra l lamina in some species. The stylets are found lateral to the buccal tube (Figure 4) and can be extended into the tube and out the mouth by protractor and retractor muscles attached to the st ylet supports. The buccal tube connects to the muscular phar yngeal bulb which in most tard igrades is lined with three rows of cuticular thickeni ngs called placoids (macroplacoids and microplacoids). The placoids are posterior to the end of the buccal tube. Figure 4. Typical buccal apparatus found in tardigrades. A. Stylet; B. Buccal tube; C. Stylet support; D. Macroplacoid; E. Microplacoid; F. Pharyngeal bulb Redrawn from Ramazzotti and Maucci (1983).

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8 Usually gonochoristic, tardigrades have a single unpaired gonad but in many cases the mode of reproduction is parthe nogenic. However, during sexual reproduction the male will deposit spermatozoa into the cloacal opening of the old cuticle while the female is simultaneously molting and laying eggs in the cuticle. Once shed the cuticle is referred to as an exuvium and fertilization ta kes place inside. Some eutardigrade eggs are freely laid and often ornamental. Both th e morphology of the eggs and the spermatozoa is important in the eutardigrades (Bertolani and Rebecchi, 1996; Guidettii and Rebecchi 1996). The phylogenetic position of Tardigrada ha s often been debated (for reviews, see Ramazzotti & Maucci, 1983; Kinchin, 1994), and morphology has primarily been used to assess the relationships among so me genera within a few fami lies of tardig rades (RenaudMornant, 1982; Kristensen, 1987; Pilat o, 1989; Pollock, 1995; Bertolani & Biserov, 1996; Jorgensen, 2000; Guidetti & Bertolani, 2001). Tardigra des have been placed along an annelid-arthropod lineage, often closel y associated with onychophorans, although there have been some arguments for placing them with a group of pseudocoelomate phyla known as aschelminthes that has since been shown to be polyphyletic (Dewel & Clark, 1973a, 1973b; Kristensen, 1991; Winnepennix et al. 1995). Current evidence suggests, however, that tardig rades, onychophorans, and arthropods form a monophyletic clade known as Panarthropoda (revi ewed in Schmidt-Rhaesa et al ., 1998). Garey et al. (1999) found close agreem ent between molecular and morphology based phylogenies that included fi ve species of tardigrades, s uggesting that the characters for the current morphological taxonomy are appropriate. Although tardigrade genera, families, orders, and classes (Table 2) are each well defined by morphological characters,

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9 the relationships among the families and among ge nera within the families have not been systematically evaluated. One exception is the family Echiniscidae for which several cladistic studies have been publishe d (Kristensen, 1987; Jorgensen, 2000). Dispersal Tardigrades are dispersed throughout three main environments: terrestrial, freshwater, and marine. Their distributions in each of the habitats may be correlated with physical environmental factors. The role of moisture, site orientation and altitude on the distribution of terrestrial tardigrade species has been widely investig ated. Oxygen availability is the limiting factor in tardigrade distributi ons in all environments. It is because of oxygen availability that terrestrial tardigrades do not inhabit de nse growing thick mosses and the reason that they are found in the top few centimeters of so il around the roots of trees, the soil must also have a degree of poros ity to facilitate movement (R amazzotti and Maucci, 1983). Those in mosses and lichens, which have b een categorized into types based on their Table 2*. Higher level Taxonomy of the Tardigrada Class Order Family Eutardigrada Parachela Calohypsibiidae Eohypsibiidae Hypsibiidae Macrobiotidae Microhypsibiidae Necopinatidae Apochela Milnesiidae Heterotardigrada Arthrotardigrada Batillipedidae Coronarctidae Halechiniscidae Renaudarctidae Stygarctidae Echiniscoidea Echiniscidae Echiniscoididae Oreellidae *(from Nelson, 2002)

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10 moisture content, need altern ating periods of wet and dry. The most studied aspect of physical environmental factors on terrestrial tardigrades is that of altitude. Many authors have reported that altitude has a definite effect on tardigrade distribution (RodriguezRoda, 1951; Nelson, 1973, 1975; Ramazzotti and Maucci, 1983; Dastych, 1985, 1987, 1988; Beasley, 1988), with most suggesting that species richness incr eases as altitude increases. Some authors (Ramazzotti and Maucci, 1983; Dastych, 1987, 1988) have even classified tardigrades based on the locality (lowland, upland, montane, etc.). However, others have reported that dist ributional patterns were not in fluenced by altitude (Kathman and Cross, 1991). Very little information exists as to the m ode of dispersal of fr eshwater and marine tardigrades since very few undergo cryptobiosis They are most lik ely distributed by way of rapid moving waters during periods of flooding, or by storm surge altering the coastal currents. Tardigrade zonation in littoral habita ts, like that of terrestrial species, is limited by the availability of oxygen. Size of the sand grain and circulation of the water may also affect zonation patterns. Both will have an effect on oxygen availability which could account for the distribution of the tardigrade s in the first few centimeters of the sand (Pollock 1975). Light availability, salinity, and temperature have been shown to affect species distributions in the marine environment (Pollock 1975). Ecological Importance Meiofauna are important in many terrestrial and aquatic ecosystems (Wilson, 1992; Palmer et al. 1997). Aquatic communities cons ist of many diverse and abundant species of benthic invertebrates (includi ng tardigrades). A typical food web often

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11 consists of nematodes, tardigrades, bacter ia, algae, rotifers, protozoa, mites, and collembolans (Kinchin 1987). One possible ro le of terrestrial ta rdigrades is in the colonizing of new habitats. Tardigrades may pl ay an important role with the ability to move into a newly formed habitat quickly, es tablishing as a pioneer species and then in turn attracting other meiofaunal groups to the habitat. The habitat then becomes suitable to colonization by macrofaunal species. Because of the size of tardigrades, limited studies on the ecological role of marine and freshwater species have been performed a nd it is difficult to assess their importance in ecological functioning. Typically, they are a major player in the meiofaunal assemblages second only to that of nematodes and rotifers. They may play a role in converting plant and animal matter into f ood for larger organisms and could aid in maintaining aeration and nutrient cycling in the sediments. They may even form distinct functional groups that exists only in a few ar eas and if lost could be a detriment to the ecosystem. There are a number of studies of community structure of terrestrial tardigrades, but few (Gaugler, 2003, unpublished thesis) focu sed on community structure of marine species. Currently there are no models th at can accurately meas ure their role in ecosystem functioning therefore, it would be impossible to assess their importance. Significance There currently is no phylogenetic hypothesi s of tardigrade evol ution that treats the group as a whole. Instead, th e tardigrade literature consis ts of a plethora of species descriptions with very few systematic studies. A molecular framework for tardigrade

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12 phylogeny would be very useful and woul d encourage a wider application of morphological and molecular studies to ta rdigrade phylogeny. For example, a recent molecular study of nemat ode phylogeny (Blaxter et al., 1998) was pivotal in encouraging new discussion and a reassessment of nemat ode phylogeny, particularly in light of the completion of the Caenorhabiditis elegans genome project. Tardigrada, as a member of Panarthropoda will likely play and important role in the future assessment and discussion of two competing theories of protostome evolution, that of Ecdysozoa (Aguinaldo et al., 1997) and Articulata (Cuvier, 1863) in the same way as onychophorans (de Rosa et al ., 1999). Tardigrades, like onychophorans (e.g. Panganiban, et al. 1997, Grenier and Carroll, 2000) will likely become more importa nt in discussions of the evolution of body plans and appendages as well. It has been noted that species or even familial identification of tardigrades can be difficult for untrained investigators. For example, specimens of Macrobiotus species have been sold commercially as Milnesium tardigradum (Garey et al., 1999). As tardigrades are used more extensively as model organisms for molecular and developmental studies, it becomes very important that specimens are properly identified.

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13 Objectives 1. The present study was designed to provi de an overall framework for tardigrade evolution utilizing morphology and mol ecular based (18S rRNA) phylogenetic hypotheses for tardigrades. Potential morphologi cal characters that ar e informative at the family level were assembled and a morphol ogy based phylogeny was determined. A data set of 18S rRNA gene sequences was assembled and a molecular based phylogeny was developed. These data were used to investigate the phyloge netic relationships among the tardigrade families and to test the relations hips among the families within each tardigrade order to determine if the families are trul y monophyletic. Finally, it has been suggested that tardigrades evolved in a marine envir onment (May 1953) and that some echiniscids adapted to freshwater and then to a terrestri al habitat. The question is asked, what was the ancestral state and how many times did the adaptations occur? 2. The purpose of this study was to inve stigate the structure (frequency, diversity, similarity) of a meiofaunal community that has an usually abundant tardigrade population using both morphological specie s identification and molecula r species iden tification. Individuals were grouped to th e lowest practical taxon and the abundance, diversity and similarity between the “morphological community” and “molecular community” was determined.

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14 Chapter 2 Specimen Acquisition and Processing Specimens were obtained from a network of tardigrade researchers and from field collections. Marine specimens were colle cted and rinsed with fresh water. The freshwater rinse puts the tard igrades into osmotic shock a nd they release their grasp on the sand grains. These samples were then ri nsed through a set of nested sieves, which have mesh openings from 1600 mm to 40mm, with seawater to both collect the individual and stop the osmotic shock (Higgins and Thie l 1988). The specimens were drained into a screw-top jar and preserved by addi ng 100% ethanol or stored at –80O C for DNA extraction. Dried samples of cryptogams were soaked in tap water in a stoppered funnel for 12-24 hours. The cryptogams were then ag itated, removed, and squeezed to drain the remaining water into the funnel. Samples were processed by sieving through nested sieves. The specimens were then sorted, st ored in screw-top ja rs, and preserved. The cryptogams were re-dried and stored in pr eviously labeled paper bags. The preserved samples were placed into a Syracuse dish and searched with a stereoscopic dissecting microscope at a magnification of 7-45X. A pipette was used to extract individual tardigrades to separate glass slides with glass cover slips (18mm circle #1 glass) for identification.

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15 Chapter 3 Morphological Analysis of Tardigrade Phylogeny Background Hennig (1979) stated that a classificati on should express the br anching (cladistic) relationships among species, re gardless of their degree of similarity or difference. Cladistic analysis is based on common ancestr y rather than overall similarity with emphasis placed on character types and the importance of those characters (Forey et al., 1992). For example, a cladistic analysis mu st determine the polarity of the character states to show which are pleisiomorphic (an cestral) and which are apomorphic (derived). Cladistic analyses utilize parsimony met hods to construct a cladogram. This method implies that the simplest answer is us ually correct or in the case of a phylogenetic tree, the shortest tree is usually correct. Thus the advantage of cladistic analysis utilizing polarized character states is that it repr esents the topology of a phylogenetic tree and can be used to infer evolutionary history of the groups being stud ied. Cladistic analyses are limited to monophyletic groups, only decendents of a particular ancestor can be included and paraphyletic groups are not allowed. When performed correctly, complications by convergent evolution, parallelis m and reversal are avoided. However, differing opinions on character state definitions between researcher s and the fact that fossil material may be needed to evaluate a particular missi ng character can be a disadvantage.

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16 The phylogenetic position of Tardigrada ha s long been debated (for reviews, see Ramazzotti & Maucci, 1983; Kinchin, 1994), and morphology has been used to assess the relationships among some genera within a few families (Renaud-Mornant, 1982; Kristensen, 1987; Pollock, 1995; Bertolani & Biserov, 1996; Jorgensen, 2000; Guidetti & Bertolani, 2001). Tardigrades have been pl aced along an annelid-a rthropod lineage, often closely associated with the onychophoran-art hropod complex, although there have been some arguments for placing them with the aschelminthes group (Dewel & Clark, 1973a, 1973b; Kristensen, 1991). Current evidence suggests, however, that tardigrades, onychophorans, and arthropods form a monophyl etic clade known as Panarthropoda (reviewed in Schmidt-Rhaesa et al., 1998). The present study was designed to assemb le and analyze potential morphological characters that are informative at the family level and to test the relationships among the families within each tardigrade order. Materials and Methods Characters A matrix (Appendix A) consisting of 50 morphological char acters (Appendix B) from current literature was scored for 15 tard igrade families (for review, see Schuster et al., 1980; Binda & Kristensen, 1987; Kriste nsen, 1987; Bertolani & Rebecchi, 1993; Schmidt-Rhaesa et al., 1998; Jorgensen, 2000; Guidetti & Bertolani, 2001). Characters common to most families but with character st ates that distinguished families from one another were chosen. Three outgroups; two within Ecdysozoa (Lor icifer & Kinorhyncha)

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17 and one outside Ecdysozoa (Gastrotricha) were included in the analysis. Ten multi-state characters were present. Data Analysis The character data matrix was cons tructed using Nexus Data Editor (NDE) version 0.5.0 (http://taxono my.zoology.gla.ac.uk/rod/NDE/nde.html). Phylogenetic Analysis Using Parsimony (PAUP*) versi on 4.6 (Swofford, 1998) was utilized for maximum parsimony analysis. Results and Discussion Morphological Phylogeny The 50% majority rule tree in Figure 6 was generated from an analysis of the morphological data set and agrees with the consensus that Eutardigrada and Heterotardigrada are distinct monophyletic si ster groups as eviden ced by a high bootstrap value of 98%. Among the eutardigrades, Eohypsibiidae was a sister group to Macrobiotidae + Hypsibiidae, though the bootstrap suppor t around 60% was minimal. Necopinatidae appears to be basal among the Parachela, which forms a monophyletic sister group to the Apochela and the most ba sal eutardigrade fam ily, Milnesiidae (Figure 5). Among the heterotardigrade families, bootst rap values indicate that all branches were moderately or highly supported (Fi gure 6). Kristensen & Higgins (1984)

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18 established the family Renaudarctidae based on the conclusion that the development of toes and adhesive discs in Halechiniscidae and Batillipedidae were derived characters and that the claw insertion in Stygarctidae was a plesiomorphic state. This assessment agreed with Renaud-Mornant’s (1982) suggestion that Stygarctidae should be the most primitive Figure 5. Maximum parsimony phylogeny of tardigrade families with mapped habitat preferences. The analysis supports the mon ophyletic origin of the class Eutardigrada and orders Parachela and Apochela along w ith the class Heterotardigrada; however, the orders within Heterotardigrada (Arthr otardigrada, Echiniscoidea) may not be monophyletic. Key to Characters: 1. Eu tardigrada: No cephalic appendages; lack of dorsal plates; differentiated placoids; do uble claws with secondary and primary branches. 2. Heterotardigrada: Cephalic appendages, digitate legs with or without claws. 3. Parachela: No cephalic papillae; double claws per leg divided into a secondary and primary branch. 4. Apochela : Six cephalic papillae; two sets of claws per leg, with unbranched primary claw separated from secondary claw; 4 6 peribuccal lamellae. Habitats: Terrestrial (T); Marine (M); or Freshwater (F). Maximum parsimony bootstrap values are shown.

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19 heterotardigrade family. The current anal ysis (Figure 5) suggests instead that Halechiniscidae is more basal than eith er Stygarctidae or Renaudarctidae and that Oreelidae is the most basal heterotardigrade. This placement supports earlier suggestions by Thulin (1928) and Marcus (1929) that Echi niscidae is derived from arthrotardigrades and that Oreellidae should be considered the most primitive heterotardgrade. The cephalic appendages that define heterotardigrade s are present in oreell ids, but the dorsal plates, characteristic of othe r heterotardigrades, are completely absent. The morphology of the various dorsal plates appears to provi de important characters in heterotardigrade phylogeny. If the absence of dorsal plates represents a secondary loss within the oreellids, then the basal position of Oreelli dae in this study may be an artifact. The orders Arthrotardigrada and Echinisc oidea have been placed as sister groups based on shared derived characters (Eibye-J acobsen, 2001). The current results (Figure 5) suggest that these orders ar e not sister groups, and it appe ars that Arthrotardigrada is paraphyletic containing some members of Echiniscoidea. Habitat Mapping Mapping of habitat preferences onto the character tree (Figure 5) suggests that tardigrades have adapted to marine environmen ts twice, to freshwater environments at least three times, and to terrestrial environmen ts twice. The placement of the terrestrial Oreellidae as the basal heterotardigrade s uggests that tardigrades were ancestrally terrestrial. However, the placement of Oreellid ae in the character tree may be an artifact caused by the secondary loss of dorsal plates.

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20 Chapter 4 Molecular Analysis of Tardigrade Phylogeny Background Molecular studies have examined the phylogenetic position of the Tardigrada (Garey et al., 1996; Giribet et al ., 1996) placing them in a clad e that includes arthropods. There appears to be a consensus that ar thropods, onycophorans and tardigrades form a monophyletic clade known as Panarthropoda (reviewed in Schmidt-Rhaesa et al., 1998). An on-going study to use protein coding gene sequences to study arthropod phylogeny utilized several tardigrades as an arth ropod outgroup (Regier & Shultz, 2001). Other molecular studies have placed tardigrades w ithin a group of molting animals that includes arthropods, priapulids, nemat odes and kinorhynchs (Aguinaldo et al., 1997; Zrzavy et al., 1998). Garey et al. (1999) found close agreement be tween molecular and morphology based phylogenies that included si x species of tardigrades, su ggesting that the characters for tardigrade morphological st udies are appropriate. This study also suggested that Heterotardigrades, the marine and terrestria l armored species, are the most basal group with the greatest number of plesiomorphic characters. Garey et al (1996) showed that tardigrades ar e closely allied to arthropods and that tardigrades, arth ropods and priapulids form a clade. Since priapulids had long been

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21 considered to be members of “aschelminths”, this explained why some tardigrade characters such as cryptobiosis, cuticular structure and the presence of a triradiate pharynx seemed to ally them to aschelmi nths while other characters such as body muscle specialization, lack of a closed ci rculatory system, and body segmentation placed tardigrades most closely with arthropods (Garey et al., 1996). A similar molecular study, published the same year by Giribet et al. (1996) also demonstrated that tardigrades are associated with arthropods. There are a number of reports where molecular and morphological data have been evaluated t ogether in studies of rotifer phylogeny, tardigrades, protostomes, urochor dates, and deuterostomes (Cameron et al., 2000; Swalla et al., 2000; Garey et al ., 1998; Garey et al., 1999; Schmidt-Rhaesa et al., 1998; Zrzavy et al., 1998; Ernissee, 1992; Giribet et al., 2001) Garey et al. (1999) demonstr ated that trees produced by the analysis of 18S rRNA gene sequences of tardigrade s are in close agreement to trees based on morphological characters (Fig. 6). This suggests that mol ecular analyses of tardigrade 18S rRNA gene sequences contain sufficient information to outline the evolutionary relationships among tardigrade orders, among tardigrade families, and even among tardigrade genera.

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22 Materials and Methods Gene Selection The near complete 18S rRNA gene sequence (~1800 b; Fig. 7) was used for the molecular analysis. Nuclear protein coding genes were consid ered and rejected for this study because of the small sample size and th e relative difficulty in obtaining both DNA Figure 6. Topology of an 18S rRNA based tree mapped with diagnostic morphological characters (Garey et al. 1999). The molecular tree is completely congruent with current morphological hypotheses of tardigrade phyloge ny. Bootstrap and confidence probability values are for NJ and MP trees are shown at each node. Key to Characters: 1. Arthropoda + Tardigrada: supraesophageal or preoral position of the frontal appendages and their neuromeres (DEWEL & DEWEL 1997). 2. Arthropoda: body with articulated exoskeleton; protocerebrum with compound eyes (NIELSEN 1995). 3. Tardig rada: connective between protocerebrum and ganglion of first pair of legs DEWEL & DEWEL 1997). 4. Heterotardigrada: Cephalic appendages, legs with digits and/or claws ( BARNES & HARRISON 1993). 5. Eutardigrada: Lack of cephalic appendages; legs with claws bu t not digits (BARNES and HARRISON 1993). 6. Apochela: With cephalic papillae and with dou ble claws with well-separated primary and secondary branches (SCHUSTER et al. 1980). 7. Parachela: Without cephalic papillae and with double claws in which primary and secondary bran ches are joined (SCHUSTER et al. 1980). 8. Macrobiotus: Claw branches with sequence: seco ndary, primary, primary, secondary; buccal tube with ventral lamina and 10 peribuccal lamellae (SCHUSTER et al. 1980). 9. Thulinia + Hypsibius: Claw branches with sequence: seconda ry, primary, secondary, primary; buccal tube without ventral lamina (SCHUSTER et al. 1980; BERTOLANI 1982). 10. Thulinia: twelve peribuccal lamellae (BERTOLANI 1982). 11. Hypsibius: peribuccal lamellae absent (SCHUSTER et al. 1980).

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23 Figure 7. This study used the 18S rRNA gene using primers to amplify the portions of each gene as shown. See text for details. and poly A+RNA from limited (e.g. size and number) specimens. The 18S rRNA gene was used based upon its recent use to infer intraphylum relationships (Blaxter et al ., 1998) and that it has a long history in determining interphylum relationships among metazoans (Field et al ., 1988). DNA Extraction Tardigrades were isolated onto indivi dual glass slides and air dried. The tardigrade cuticle was then mechanically macerated using dental probes previously cleaned with DNA-Away (Molecular BioProduc ts, Inc., San Diego, CA) and rinsed in deionized water. A 10 l solution containing 5mg/ml of Proteinase K was added to the macerated cuticle and the sample was tr eated to 3 freeze/thaw cycles at -20 oC. Proteinase K cleaves peptide bonds and is used for the removal of DNases and RNases during DNA and RNA isolation and has been show n to improve the efficiency of cloning in PCR products (Crowe et al ., 1991). The extraction mixtur e was then transferred to a 200 l centrifuge tube and incubated at 55 oC for one hour, 5 minutes at 95 oC and then centrifuged at 2000 x g for 5 minutes. Ten l of the supernatant was then used as template DNA.

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24 Polymerase Chain Reaction PCR amplification of template DNA was ca rried out using primers specific for the 18S rRNA gene (18S4 5'-GCTTGTC TCAAAGATTAAGCC-3' and 18S5 5'ACCATACTCCCCCCGGAACC-3'). PCR reacti ons were performed in 200 l tubes using a Biometra TRIO thermocycler (WhatmanBiometra, Gttingen, Germany). The reaction cycle consisted of an initi al denaturing step of 30 sec at 94 oC followed by 35 cycles of 30 seconds denaturing at 94 oC, 30 seconds annealing at 45-55 oC, and 60-120 seconds extension at 72 oC. The PCR reactions consisted of 1X final concentration of 10X PCR buffer (Enzypol, Denver, CO), 2 mM final concentration of magnesium chloride, 0.1 M final concentration of each primer, 0.25 mM final concentration for each of dATP, dCTP, dTTP, and dGTP, 10 l of template DNA and 1 unit of EnzyPlus 2000 Taq polymerase (Enzypol, Denver, CO) in a final volume of 50 l. Cloning PCR product of 18S rDNA amplified from some of the samples was cloned using the TOPO TA cloning kit for sequencing (In vitrogen Corp., San Diego, CA) following the manufacturer’s instructions. Transformed ce lls were plated and incubated overnight at 370C on Luria-Bertani (LB) agar containing 100g/mL ampicillin and 50g/mL X-gal (5-Bromo-4-chloro-3-indolyl -Dgalactoside). Colonies were picked and cultured in 96 well microtiter plates for 24 hours. Plasmid extraction from the bacteria was performed using the Eppendorf Perfectprep Plasmid Isol ation Kit and then quantified using 0.9% agarose gel electrophoresis. All colonies grown overnight fo r isolation of plasmid DNA were preserved in 50% gly cerol and stored at –80 oC

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25 Sequencing Template DNA (cloned and direct PCR product) was cycle sequenced using the QuickStart sequencing kit (BeckmanCoulter, La Jolla, CA). The r eaction mix contained between 50 and 100ng of template, 2 l se quencing reaction mix (SRM), 1l 3.2 M sequencing primer, and water to bring the to tal reaction volume to 10 l. All products (clones and PCR) for analys is were sequenced using the 18S4c and 18S2c primer. Sequencing reactions were amplified in the Bi ometra TRIO thermocycler. A preheat step consisting of only water and te mplate DNA was performed at 96 0C followed by a return to room temperature. The SRM and primers were added and the reaction was cycled as follows: an initial denature at 96 oC for 1 minute; cycled 25 times through a 96 oC, denature step for 15 seconds, a 50 oC annealing step for 30 sec onds and an extension step at 60 oC for one minute. Finally, a 60 oC extension step for 7 mi nutes was performed and a final hold at 4 oC. The cycle sequencing product was purified following the manufacturer’s instru ctions and analyzed using a CE Q 8000 Genetic Analysis system (BeckmanCoulter, La Jolla, CA). Alignments and Sequence Analysis Sequences were visually checked and co rrected for ambiguous bases (N’s). Data sets from individual tardigra des were aligned according to a secondary structure model (Neefs et a l. 1993) using DCSE (De Rijk and De W achter 1993). Phylogenetic analysis of alignments was performed using MEGA ve rsion 2.1 (Kumar, et al. 2001) to produce neighbor-joining trees and Phylogenetic Anal ysis Using Parsimony (PAUP*) version 4.6 (Swofford, 1998) was utilized for Maximum Pa rsimony analysis. A variety of arthropod

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26 species were used as outgroups. Trees pr oduced were evaluated by bootstrap analysis (Hillis & Bull 1993). Results & Discussion Molecular Phylogeny Near complete 18S rRNA sequences were amplified and aligned with previously published data (Table 3). Table 3. List of species, sequence le ngth, origin, and sequence reference. Species Length Accesion Reference Arthropoda Artemia salina 2002 X01723 Nelles et al ., 1984 Meloe proscarabaeus 1934 X77786 Chalwatzis et al., 1995 Okanagana utahensis 1918 U06478 Campbell et al ., 1995 Tenebrio molitor 2083 X07801 Hendriks et al., 1988 Panulirus argus 1872 U19182 Trapido-Rosenthal et al ., 1994 Mollusca Placopecten magellanicus 1814 X53899 Rice 1990 Priapulida Priapulus caudatus 1814 X80234 Winnepenninckx et al ., 1995 Tardigrada Echiniscus viridissimus 1824 AF056024 Garey et al., 1996 Halechiniscus remanei 827 AY582118 Jorgensen & Kristensen, 2004 Halobiotus stenostomus 1783 AY582121 Jorgensen & Kristensen, 2004 Macrobiotus hufelandi 1808 X81442 Giribet et al ., 1996 Macrobiotus tonolli 1735 U32393 Garey et al., 1996 Milnesium tardigradum 1844 U49909 Aguinaldo et al., 1997 Milnesium tardigradum 1777 AY582120 Jorgensen & Kristensen, 2004 Pseudechiniscus islandicus 1820 AY582119 Jorgensen & Kristensen, 2004 Ramazzottius oberhauseri 1771 AY582122 Jorgensen & Kristensen, 2004 Thulinius stephaniae 1686 AF056023 Garey et al., 1996 Batillipes mirus 1432 Present study Present study Hypsibius dujardini 1521 Present study Present study Calohypsibius schusteri 1384 Present study Present study Ramazzottius oberhauseri 1674 Present study Present study Richtersius coronifer 1726 Present study Present study

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27 The NJ and MP trees produced from 18S rRNA gene sequences (Figure 8) were congruent with one another and support the morphological char acter analysis in indicating that Eutardigrada and Heterotardigrada are each monophyletic as evidenced by bootstrap values of 89 and 84, respectively. Figure 8. Tardigrade phylogeny based on l8 S rRNA gene sequences. Similar trees were recovered for maximum parsimony and maximu m likelihood analyses. Branch lengths are drawn to scale (substitutions/site). The numb ers above each fork are bootstrap values for the NJ tree. *Specimens were not identified to speci es in the original study, but have since been verified.

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28 The 18S rRNA gene sequences from Calohypsibius schusteri Hypsibius dujardini Richtersius coronifer and Ramazzotius oberhouseri adds 4 additional genera to the previously published eutardigrade data set. Within eutardigrades the monophyly of Parachela is well supported. Bootstrap values for all branches were high with the lowest value of 90% for the Hypsibiidae + Macrobiotidae node. The Calohypsibius sequence indicates that Calohypsibiidae is a sister group to Hypsibiidae (Figure 8). However, in the morphological tree, the relationship between Calohypsibiidae and Hypsibiidae is unresolved (Figure 6). There is more stat istical support for rele vant nodes within Eutardigrada in the molecu lar results (82% 100% boot strap values) than in the morphological results (54% 64% bootstrap va lues). This analysis agrees with the findings of Jorgensen and Kristensen (2004) that the eutardigrades are monophyletic and it is consistent with the morphological analys is in Figure 5. However, the taxon sampling among the Macrobiotidae family is not exha ustive and some questions about the monophyly of this line s till exist. A separate analysis of the Macrobiotidae family by Guidetti et al (2005) used both morphological characters and molecular data from the Cytochrome c Oxidase subunit I gene. Cytochrome c Oxidase subunit I sequences have been shown to be useful in resolving phylogenetic relationships among closely-related taxa among different phyla (Avise 1994, 2000; Brown et al. 1994; Lunt et al. 1996; Hebert et al. 2003a, b). Their findings suggest that Macrobiotidae is not monophyletic and that the subfamily Murrayinae should be elevated to family leve l Murrayidae. Nearly 40% of the Parachela genera are currently within the family Macrobi otidae and this is clear ly an area that needs further study in order to determine an accurate phylogeny.

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29 Within heterotardigrades, the monophyly of Arthrotardigrada and Echiniscoidea is well supported with a bootst rap value of 84%. There is more statistical support for relevant nodes in the molecular results ( 90% 100% bootstrap va lues) than in the morphological tree (60% 64% bootstrap valu es). The 18S rRNA gene sequence from Batillipes mirus and Halechiniscus remanae (Arthrotardigrada) adds an additional heterotardigrade order and two new families (Batillipedidae, Halech iniscidae) to the previously published molecular data set. The addition of Pseudechiniscus islandicus adds an additional genus within the Echinisc idae family to the dataset. The molecular data suggests that hetero tardigrades and eu tardigrades are both monophyletic. Habitat Mapping As seen in the morphological dataset, mapping of habitat preferences onto the gene tree (Figure 9A) also sugge sts that tardigrades have adap ted to marine environments twice, to freshwater environments once, a nd to terrestrial environments twice. The placement of B. mirus and H. remanaei as the basal heterotardigrade suggest that tardigrades were ancestrally marine. By ma pping the habitats preferences on the tree we can trace the number of evolutionary events th at occur. If terrestrial tardigrades are considered to be ancestral stat e then there are four events th at occur (Figure 9B) while if we assume that the marine tardigrades are th e ancestral tardigrades then there are five events that occur (Figure 9C). This suggest s that based on the number of evolutionary changes, terrestrial tardigrades are the ancestral tardigrades. However, more extensive taxon sampling will be required to test this hypothesis.

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30 Figure 9. Tardigrade molecular topology from Fig. 8 with mapped habitat preference. Habitats: Terrestrial (T ); Marine (M ); or Freshwater (F ).

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31 Chapter 5 Meiofaunal Abundance, diversity and Similarity in a Uniquely Rich Tardigrade Community. Background Meiofauna are distributed worldwide a nd include representa tives of 22 of the metazoan phyla (Coull, 1999, 1988). They are f ound in terrestrial, freshwater and marine habitats and are classified by their sma ll size, generally 50 500 m. Ecologically, meiofauna function as food sources for higher tr ophic levels and play an important role in the biomineralization of organic matter (Hummon, 1987; Battigelli & Berch, 1993; Coull, 1999). The assemblages can be linked to se diment characteristics (particle size) and bacterial production (H iggins, 1988; Vanreusel et al., 1995; Schratzberger et al., 2000) and are often reported in terms of abundan ce, distribution and di versity (eg. Hummon, 1987; Trett et al ., 2000, Battigelli & Berch, 2004). Meiofaunal assemblages in marine habita ts have been extensively studied and focused on areas from intertidal mud flats and littoral zones (Coull & Wells, 1981; Stoffels et al ., 2003) to the bathyal and abyssal depths of the deep sea (Ingole et al ., 2000; Tselepides et al ., 2004). Vertical and horizontal zonation is affected by an anaerobic-aerobic boundary layer in sediment, increasing dept h and salinity gradients. Typically, the highest abundances are found in intertidal zone s and decrease as the depth

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32 increases. Salinity, food availability, sedime nt grain size, and tidal exposure can affect dispersal (Findlay, 1981) which is often patchy with densities changi ng over a distance of just a few centimeters (Schratzberger et al., 2000). Surveys on marine tardigrade distributi ons are primarily descriptions of new species and taxonomical reviews (De Zi o Grimaldi & D’Addabbo, 2001). Studies investigating the ecology of marine tardig rades most often do not report information on their abundance and ecology within the cont ext of the meiofaunal community. A few exceptions are from ecological studies from th e Mediterranean Sea (for review see, De Zio Grimaldi & D’Addabbo, 2001), psammolittoral tardigrades from North Carolina (Lindgren, 1971), interstitial ta rdigrades from the Pacific coast of the U.S. and the Galapagos (McKirdy, 1976; Pollock, 1989) and the Faroe Bank (Hansen et al ., 2001). An unpublished master’s thesis (Gaugler, 2002) investigated the distribution pattern of meiofaunal at Huntington Beach, SC with particular emphasis on tardigrades. Numerous reports have investigated me iofaunal assemblages and used them as a measure of the health of the e nvironment (for example, Austen et al ., 1989; Schratzberger et al. 2000; Ingole et al ., 2000; Schratzberger & Jennings, 2002). The most difficult and prohibitive aspects of these investigations are the high cost of sample processing, difficulty associated with species identif ications and the need for broad taxonomic expertise (Warwick, 1988), an area of st udy that has declined in recent years. Traditional sorting techniques have in volved osmotic shocks and freshwater rinses, Ludox gel isolation, air bubbling, and hand sorting. These methods are time consuming and in some cases can result in damage to the studied organisms (Higgins, 1988). Molecular techniques applied to sedi ment extracts may provide a more cost

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33 effective and less time consuming method of identifying the structure of a meiofaunal community. Microbial ecologists often experience similar difficulty when attempting to isolate and culture microbes from e nvironmental samples (Stephen et al ., 1996) and those that have been cultured represent only a fracti on of the estimated species (Wintzingerode et al., 1997). Techniques develope d to extract DNA from sedi ment samples have allowed microbial ecologists to analyze the speci es composition of unculturable microbes (Kennedy and Gewin, 1997). The techniques de veloped for microbial ecology should be applicable to meiofaunal studies and similar investigations have yielded promising yet mixed results (Boenigk et al. 2005; Savin et al. 2004; Blaxter et al. 2003; Hamilton 2003). Boengk et al (2005) reported high community diversity with morphological identifications and molecular identification of plankton diversity in the Bay of Fundy. However, even though the community diversity was high, the similari ty between the two was low with few species common to both the morphological and molecular analysis (mainly diatom, dinoflagellates, etc.). Savin et al. (2004) isolated flagellates from freshwater sediments and soils to investig ate the diversity using rRNA sequence data. They were able to determine groups and iden tify the groups to closely related taxa. Blaxter et al (2003) found a high diversity when targeting tardigrade species and utilizing DNA extracts from sediment and moss samples. Hamilton (2003), like Boengk et al. (2005) found discrepancies between the nu mbers of taxonomical units recovered from sequencing sediment extracts and the traditional methods of performing manual identification and counts. Hamilton (2003) also demonstrated that increasing the size of

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34 the DNA fragment (~230 – 400bp) also increased the percentage of OTU’s that could be identified to a specific taxon from 15% to 70% with bootstrap values to support them. Materials and Methods Sample Collection Refer to Chapter 2 for general methods on sample collecting and processing. DNA Sediment Extraction A modification of Hempstead’s protocol (Hempstead et al ., 1990) for DNA extraction was used to obtain meiofauna l DNA from marine sediment collected on Dauphin Island, Alabama. Sediment samples we re collected from a small sand bar in the salt marsh on the southwest side of the airport runway (30o 15’ 26.11” N/88o 07’ 27.14” W) (Figure 10). Figure 10. Location of sediment collection s ite on Daupin Island, AL (15M Resolution). One volume (about 1 mL) of homogenization buffer (3.5% SDS in 1M Tris, pH 8.0, and 100mM EDTA) was added to a 2 mL centrifuge tube containing the sediment sample.

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35 The samples were then homogenized usi ng pre-cleaned Teflon tipped pestles and centrifuged to pellet the sediment. The supe rnatant was pipetted to a 1.5 mL centrifuge tube and an equal volume of phenol (pH 7.9) wa s added to each of the tubes. The tubes were mixed gently for 5 minutes and then centrifuged for 5 minutes at 14,000 rpm. The top aqueous layer from the solution was transf erred to a new 1.5mL tube and the previous steps were repeated: one more time using phe nol (pH 7.9), twice using a 1:1 solution of phenol (pH 7.9): chloroform-isoamyl alcohol (24 parts chloroform to 1 part isoamyl alcohol), and twice with the chloroform-isoamyl solution. The DNA in the final aqueous layer was transferred to a new tube an d was precipitated for 24 hours at –20 oC with 2 volumes of 100% ethanol and a 0.1 volume of 3M sodium acetate (pH 6.0). The precipitated DNA was pelleted by centrifugation at 14,000 rp m for 15 minutes, washed with 70% ethanol and suspended in 100 l of deionized water. Polymerase Chain Reaction PCR amplification of template DNA wa s carried out using universal primers specific for the 18S rRNA gene (18S4 5'-GCTTGTCTCAAAGATTAAGCC-3' and 18S5 5'-ACCATACTCCCCCCGGAACC-3') PCR reactions were performed in 200 l tubes using a Biometra TRIO thermocycler (WhatmanBiometra, Gttingen, Germany). The reaction cycle consisted of an initi al denaturing step of 30 sec at 94 oC followed by 35 cycles of 30 seconds denaturing at 94 oC, 30 seconds annealing at 55 oC, and 60-120 seconds extension at 72 oC. The PCR reactions consisted of 1X final concentration of 10X PCR buffer (Enzypol, Denver, CO), 2 mM final concentration of magnesium chloride, 0.1 M final concentration of each primer, 0.25 mM final concentration for

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36 each of dATP, dCTP, dTTP, and dGTP, 10 l of template DNA and 1 unit of EnzyPlus 2000 Taq polymerase (Enzypol, Denver, CO) in a final volume of 50 l. Cloning PCR product of was cloned using the TOPO TA cloning kit for sequencing (Invitrogen Corp., San Diego, CA) followi ng the manufacturer’s instructions. Transformed cells were plated and incubated overnight at 37 oC on Luria-Bertani (LB) agar containing 100g/mL ampicillin and 50 g/mL X-gal (5-Bromo-4-chloro-3-indolyl -Dgalactoside). Colonies were picked and cultured in 96 well microtiter plates for 24 hours. Plasmid extraction from the bact eria was performed using the Eppendorf Perfectprep Plasmid Isolation Kit and th en quantified using 0.9% agarose gel electrophoresis. All colonies grown overnight for isolation of plasmid DNA were preserved in 50% glycerol and stored at –80 oC Sequencing Template DNA (cloned and direct PCR product) was cycle sequenced using the QuickStart sequencing kit (BeckmanCoulter, La Jolla, CA). The r eaction mix contained between 50 and 100ng of template, 2 l se quencing reaction mix (SRM), 1l 3.2 M sequencing primer, and water to bring the to tal reaction volume to 10 l. All products (clones and PCR) for anal ysis were sequenced using the 18S4 and 18S5 primer. Sequencing reactions were amplified in the Bi ometra TRIO thermocycler. A preheat step consisting of only water and te mplate DNA was performed at 96 oC followed by a return to room temperature. The SRM and primers were added and the reaction was cycled as follows: an initial denature at 96 oC for 1 minute; cycled 25 times through a 96 oC,

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37 denature step for 15 seconds, a 50 oC annealing step for 30 sec onds and an extension step at 60 oC for one minute. Finally, a 60 oC extension step for 7 mi nutes was performed and a final hold at 4 oC. The cycle sequencing product was purified following the manufacturer’s instru ctions and analyzed using a CE Q 8000 Genetic Analysis system (BeckmanCoulter, La Jolla, CA). Alignments and Sequence Analysis Sequences were visually checked and corrected for ambiguous bases (N’s) and aligned using ClustalX (Thompson, et al 1997). Phylogenetic anal ysis of the alignment was performed using MEGA version 2.1 (Kumar et al ., 2001) to produce neighborjoining trees evaluated by bootstrap analysis (Hillis & Bull, 1993). Operational taxonomic units (OTUs) were assigned to sequences from the tree containing all 126 sequences based on the number of differen ces and the topology of the tree. Assigned OTUs were given to sequences that grouped together as a clade and had less than 5 differences. Sequence misalignments were manually corrected. Sediment sorting and sp ecimen identification Meiofaunal samples preserved in 95% ethanol with Rose Bengal stain were manually sorted and counted using a Meiji diss ecting scope. Type of species present and frequency of individuals were keyed to the mo st practical taxonomic level (ie. tardigrade, nematode, and ostracod). Data Analysis One sequence from each of the assigned OTUs was randomly selected and added to a data set of reference sequences. Th is dataset was aligne d with ClustalX. A

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38 phylogenetic tree utilizing the neighbor-j oining method and the Kimura 2-parameter distance model was generated. The OTUs we re identified and numbers of individual clones per OTU were scored. A rarefaction curve and species diversity indices were plotted using BioDiversity Pro (http://www.sams.ac.uk/). Species di versity was calculate d using the ShannonWiener index (H’= pi log pi) and Simpson’s index (C= 1pi 2), where pi is the number of individuals of a species divided by th e total number of individuals. Community similarity was calculated usi ng the Jaccard coefficient (CCj= c/s1 + s2 – c); where c is the number of species shared between the communities and S1 and S2 are the number of species in community 1 and 2 respec tively; and proportional similarity. The two communities compared in this study are defined as 1) the OTUs from the sequence analysis and 2) the manua l counts from preserved samples. Results and Discussion The clone library from a sediment sample with a uniquely high concentration of tardigrades at Dauphin Island, Alabama yielde d 126 sequences that were used for the analyses. Seventeen OTUs were assigned from the neighbor-joining tree (Figure 12). The tree (Figure 13) generated from a re ference data set and unique sequences representing each of the OTUs identified twelve groups (Table 4). Hamilton (2003, unpublished thesis) showed that specific sequen ces of meiofaunal data could be identified from a reference alignment. The frequencie s of individual sequences from each of the 12 groups identified here ar e reported in Table 4.

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39Table 4. Sequence groups and the number of sequences per group. These data were used to calculate Shannon diversity, Simpson’s Diversity Jaccard’s Similarity, and Proportional Similarity. OTU # Sequence # Closest Genus Common Name Frequency 1 S1140 Echiniscus Tardigrade 27 2 B1-10 Ballanus Barnacle 1 3 S240 Plectus Nematode 7 4 66-18S4 Bugula Bryozoan 1 5 6718S4 Bugula Bryozoan 1 6 561864 Electra Bryozoan 28 7 40-18S4 Vaccinium Plant 1 8 6018S4 Vaccinium Plant 1 9 4418S4 Vaccinium Plant 12 10 S381 Stenostomum Flatworm 3 11 B1-9 Wilsonema Nematode 1 12 S2B7 Callinectes Decapod 1 13 S2B6 Aurila Ostracod 6 14 S39 Caligus Copepod 5 15 39-18S4 Callinectes Decapod 1 16 S215 Daptonema Nematode 8 17 B1-8 Enoplus Nematode 1 18 B1-5 Chaetonotus Gastrotrich 8 19 S341 Abyssothyris Brachiopod 6 20 S1B46 Brachionus Rotifer 8 Total # of groups: 20 Total # of individuals: 126 Frequency data of individual OTUs (Tab le 4) and morphological groups (Table 5) was used to calculate a rarefaction curve to test for saturation of the identified species Figure 11. Rarefaction curve calculated for molecular OTUs and morphological groups

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40 (Figure 11). The rarefaction curve shows that molecular OTUs reached a saturation point around 100 individual sequences wh ile the morphological groups reached a saturation point at approximately 70 individual s. These results suggest that the sample size was large enough to adequately assess the communities.

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41

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42 Figure 12 (Continued)

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43 Figure 12 (Conitinued)

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44 Figure 12 (Continued) Figure 12. Neighbor-joining phylogenetic tree based on the number of differences between sequences and complete deletion of gaps of all 126 sequences. Groups notated to the right of each group consist of sequences with no more than 5 differences between them. The scale bar on the last page of the tree indicates the number of differences per length of the bar.

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45

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46 Figure 13 (Continued) Figure 13. Neighbor-joining phylogenetic tree of the unique OTU sequences and a reference data set. The phylogenetic tree was created based on the Kimura 2-parameter distance method and complete deletion of gaps.

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47 Species groups from manual sorting we re determined and the frequencies of individuals present are reported in Table 5. Ten groups were found. Tardigrades were the dominant species comprising nearly 35% of the population. Nematodes (18%), polychaetes (12%), rotifers (10%), diatoms (9%) and ostracods (8%) were the other main taxa identified. There is a striking contrast in the groups represented when compared to the molecular data set. Bryozoans, which were absent in the manual counts, and tardig rades were the most dominant groups in the molecular data set. They represented 24% and 21% respectively of the total sequences while sequences from polychaetes and diatoms are absent from the molecular data set. The presence of the fourteen sequences identified as Vaccinium an asterid plant was probably due to the presence of pollen in the sediment sample. The Frequency of individuals from the molecular OTUs in Ta ble 4 was lumped into the same species groups identified in Table 5 for the analysis Bryozoans and asterids were omitted from the analyses because because they were not included in the manual sorting efforts and their presence in and the sequences isolated were a result of contamination from Table 5. Morphological and sequence group (OTUs) frequency of individuals. These data, with noted omissions (*), were used to calculate Shannon diversity, Simpson’s Diversity, Jaccard’s Similarity and Stander’s Similarity. Species Morphological OTUs Tardigrade 128 27 Nematode 66 17 Barnacle 0 1 Polycheate 44 0 Flatworm 0 3 Rotifer 38 8 Diatoms* 34 0 Ostracod 32 6 Copepod 10 5 Gastrotrich 6 8 Molluscs 3 0 Crustacea 2 1 Brachiopod 0 6 Bryozoan* 0 30 Plant ( Vaccinium )* 0 14 Total # groups: 15 Total #: 363 Total #: 126

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48 fragments of bryozoan and pollen from the aste rids. The diatoms are not metazoan and were therefore omitted from the analyzed data. Species Diversity High species diversity is indicative of a highly complex community and is one expression of the community structure. A seri es of populations that are equally abundant will have high diversity indices, while thos e that have few dominant species and many rare species will have low diversity indices. Two diversity indices were calculated to focus on both species richness (number of species) and species evenness (number of individuals among the species). Shannon index (H ’) is sensitive to the presence of rare species while Simpson’s index (C) is more se nsitive to changes in species richness and species evenness. The results based on the data from table 5 show that the diversity among the molecular OTUs is higher than the morphol ogical groups (Table 6). The number of species present in OTUs (n=10) and morphological groups (n=9) differs by a count of 1. The distribution of individuals among the OTUs (mean = 6.83) are distributed more equitably and ranged from 0 -27 while the morphological groups (mean = 27.42) ranged from 0 128. Community Similarity Similarity between communities has often been questioned by ecologists. A measure of the similarity between two areas can provide an idea of the success of the Table 6. Shannon (Hmax) and Simpson’s (C) diversity indices for molecular OTU data and species groups Index OTUs Species groups Shannon ( H’) 0.837 0.733 Simpson’s ( C ) 0.231 0.176

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49 communities. Two methods of calculating comm unity similarity are Jaccard’s coefficient and Stander’s coefficient. Jaccard’s coeffi cient does not take into account relative distribution, but indicates the percentage of species shared between the two communities while Stander’s is a function of the number of species shared and their relative distributions. Both indices will range from 0, when no shared species are found, to 1.0, when all species are shared and have the same relative abundance. For this study, the molecular OTUs and the morphological groups were each treated as a separate community. There is a noticeable difference in the indices for the two community similarity coefficients. Jaccard’s coefficient which simply accounts for the number of taxa shared between the two communities is 58.3% (Table 7), a decrease of almost 40 points when compared to Stander’s coefficient of 98.6% similarity which is a function of the number of species shared and their relative distributions. The preserved samples contain one less species group, but over 35% of the individuals belonged to one species, Batillipes mirus and the second most abundant being nematodes at 10% Similar findings were seen in the OTUs where tardigrades were the dominant group representing 33% of the community followed by nematodes at 21%. The molecular and phylogenetic methods used here to assess a community of marine tardigrades were successful in di scriminating between the meiofaunal groups (OTUs). The results indicate th at rarefaction curves can reac h saturation and that groups can be identified from small sample sizes. These results can then be used to compare Table 7. Jaccard’s and Sta nder’s community similarity indices for molecular OTU community compared to the manual count community. Community Index Similarity Jaccard’s 0.583 Stander’s 0.986

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50 communities utilizing species diversity, comm unity similarity and other comparative analyses. The capability to identify specific gr oups utilizing gene sequences may lead to more detailed surveys of meiofaunal communities by lessening the burden on the researcher attempting to identify indi viduals through traditional techniques. Tardigrades from the family Batillipedidae ar e a littoral or sublittoral species with a few records from bathyal depths (Hansen et al ., 2001). The findings presented here are from a unique population of meio fauna containing species of Batillipes in an intertidal salt marsh. The abundance of Batillipes from this site is unusually high often with counts nearing 1000 individuals per 10 cm2 core samples (Romano, personal communication) compared to most samples which typically range from zero to several hundred individuals per 10 cm2 (e.g. McGinty & Higgins, 1968; D’ Addabbo Gallo et al., 1999). This abundance may be a result of the pres ence of a sand bar, which is unique when compared to a typical salt marsh. The intera ction between the grain size of the sand bar sediment providing the necessary habitat for a population of marine tardigrades and the minimal tidal influence associated with this ar ea may be high enough to facilitate nutrient exchange. Also, the highly dense population of tardigrades may not be unique and could simply be a result of a lack of investigati ons looking for tardigrades in intertidal salt marshes.

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51 Chapter 6 Summary Morphological Analysis of Tardigrade Phylogeny The morphological phylogeny developed here suggests that the class Eutardigrada and Heterotardigrada are both monophyletic The order Parach ela is a monophyletic sistergroup to Apochela within the eutard igrades while the order Parachela is a monophyletic sister group to the Apochela The phylogeny also suggests that the heterotardigrade orders Arth rotardigrada and Echiniscoid ea are not sister groups and it appears that Arthrotardigrada is paraphyletic containing some members of Echiniscoidea. The placement of the terrestrial Oreellidae as the basal heterotardigrade suggests that tardigrades were ancestrally terrestrial. Ho wever, the placement of Oreellidae may be an artifact due to the secondary loss of dorsal plates. Molecular Analysis of Tardigrade Phylogeny The molecular phylogeny is congruent with the morphological phylogeny in indicating that the class Eutardigrada and Heterotardigrada are each monophyletic. The monophyly of the order Parachela is well suppor ted and the molecular data showed high support for the relationship between the fam ilies. The monophyly of the Macrobiotidae family remains in question. The molecular phylogeny, in contrast to the morphological

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52 phylogeny, showed that the order Arthrotard igrada and Echiniscodea are monophyletic groups. Mapping of the habitat preferences sugg ests that terrestrial tardigrades are the ancestral state. However, more extensive taxon sampling will be required to test this hypothesis. Meiofaunal Abundance, diversity and Simila rity in a Uniquely Rich Tardigrade Community The molecular and phylogenetic methods us ed here to assess a community of marine tardigrades were successful in di scriminating between the meiofaunal groups (OTUs) identifiying 17 OTUs in contrast to the 12 morphological gr oups identified from the manual sorting efforts. Rarefaction curves suggest that the sample sizes were large enough to adequately assess the communities. The two communities were very similar but the OTUs had a higher diversity and eveness than the morphological groups. The dominant species in both communities was tardigrades.

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79 Ohtaka, A and H Morino. 1986. Seasonal chan ges in the epiphytic animals of the Potamogeton malaianus in Lake Kita-ura, with special reference to Oligochaetes. Ja. J. Limnol. 14(1):63-75. Packard, AS. 1873. Discovery of a Tardigrade. Am. Nat. 7:740-742. Panganiban G, Irvine SM, Lowe C, Roehl H, Corley LS, Sherbon B, Grenier JK, Fallon JF, Kimble J, Walker M, Wray GA, Swa lla BJ, Martindale MQ, Carroll SB. 1997. The origin and evolution of animal appendages, Proc. Nat. Acad. Sci. U.S.A. 94: 5162-5166. Peterson, CH. 1991. Intertidal zonation of marine invertebrates in sand and mud. American Scientist 79:236-249. Pilato, G. 1969. Evoluzione e nuova sistemazione degli Eutardigrada. Bollettino di Zoologia 36:327–345. Pilato, G. 1972. Structure, intraspecific vari ability and systematic value of the buccal armature of Eutardigrades. Z. Zool. Syst. Evolut-forsch 10:65-78. Pilato, G. 1973. Redescription of Haplomacrobiotus hermosillensis May, 1948, and consideration on the genus Haplomacrobiotus (Eutardigrada). Z. Zool. Syst. Evolutforsch. 11:283-286. Pilato, G. 1974. On the taxonomic criteria of the Eutardigrada. In: Higgins, RP (ed). Proceedings of the first international sympos ium on tardigrades. Memorie dell'Istituto di Idrobiologia 32: 277-303. Pilato, G. 1974. Struttura dell'armatura boccale di alcune specie di Isohypsibius (Eutardigrada). Animalia 1:43–58. Pilato, G. 1979. Correlations be tween cryptobiosis a nd other biological characteristics in some soil animals. Boll. Zool. 46:319-332. Pilato, G. 1987. Revision of the genus Diphascon Plate, 1889, with remarks on the subfamily Itaquasconinae (Eutardigrada, Hyps ibiidae). In: R Bertol ani (ed.) Biology of Tardigrades, Selected Symposia and M onographs U.Z.I., 1, Mucchi, Modena:339-372. Pilato, G. 1989. Phylogenesis and systematic arrangement of the family Calohypsibiidae Pilato, G. & Binda, M.G. 1989. Richtersius nuove nome generico in sostituzione di Richtersia Pilato e Binda, 1987 (Eutardigrada). Animalia 16:147–148. Pilato, 1969 (Eutardigrada). Z. Z ool. Syst. Evolut-forsch. 27:8-13. Pilato, G. 1992. Mixibius, nuovo genere di Hypsibiidae (Eutardigrada). Animalia 19(1/3):121-125.

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80 Pilato, G. 1993. Nuove considerazioni su lla posizione sistematica del genere Haplomacrobiotus May, 1948 (Eutardigrada). Animalia 20(1/3):13-21. Pilato, Giovanni, and Clark W. Beasley. 1987. Haplohexapodibius seductor n. gen. n. sp. (Eutardigrada, Calohypsibiidae) with remarks on the systematic position of the new genus. Animalia, Catania 14:65-71. Pilato, G, R Bertolani, a nd MG Binda. 1982. Sudio degli Isohypsibius del gruppo elegans (Eutardigrada, Hypsibiidae) con descrizione di due nuove specie. Animalia 9(1/3:185198. Pilato, G and MG Binda. 1997/98. A comparison of Diphascon (D.) alpinum Murray, 1906, D. (D.) chilenense Plate, 1889, and D. (D.) pingue Marcus, 1936 (Tardigrada), and description of a new sp ecies. Zool. Anz. 236:181-185. Pilato, G and MG Binda. 1998. Two new species of Diphascon (Eutardigrada) from New South Wales, Austalia. New Zeal and Journal of Zoology 25:171-174. Pilato, G and MG Binda. 1999. Three new species of Diphascon of the pingue group (Eutardigrada, Hypsibiidae) from Antarctic a and the Subantarctic Archipelagos. Not published yet. Pilato, G. and MG Binda. 2001. Biogeography an d Limno-terrestrial Tardigrades: Are They Truly Incompatible Binomials? Zoologischer Anzei ger 240(3-4):511-516. Pilato, G and L Rebecchi. 1992. Ramazzottius semisculptus nuova specie di Hypsibiidae (Eutardigrada). Animalia 19(1/3):227-234. Pollock, LW. 1970. Distribution and dynamics of interstitial Tardigrada at Woods Hole, Massachusetts, U.S.A. Ophelia 7(2):145-166. Pollock, LW. 1970. Reproductive anatomy of some marine Heterotardigrada. Trans. Am. Microsc. Soc. 89(2):308-306. Pollock, LW. 1971. On some British marine Ta rdigrada, including two new species of Batillipes J. Mar. Biol. Ass. U.K. 51:93-103. Pollock, LW. 1975. Tardigrada. In: Marine flor a and fauna of the northeastern United States. NOAA technical repor ts NMFS Circ-394:1-27. Pollock, LW. 1977. A tabular key to the species of marine Heterotardigrada. Proceedings of the II International Sy mposium on Tardigrada Krakow, Poland, July 28-30, 1977. Pollock, LW. 1983. A closer look at some ma rine Heterotardigrada. 1. The morphology and taxonomy of Orzeliscus. Proceedings of the third intern ational symposium on

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82 Ramlv, H. and P Westh. 2001. Cryptobi osis in the eutardigrade Adorybiotus (Richtersius) coronifer : tolerance to alcohols, temperature and de novo protein synthesis. Zoologischer Anzeige r 240(3-4):517-523. Ray, GC. 1991. Coastal-zone biodiversit y patterns. BioScience 41(7):490-498. Rebecchi, L. 1997. Ultrastructure study of spermatogenesis and the testicular and spermathecal spermatozoon of the gonochoristic tardigrade Xerobi otus pseudohufelandi (Eutardigrade, Macrobiotidae). Journal of Morphology 234:11-24. Rebecchi, L. 2001. The Spermatozoon in Tardigrades: Evolution and Relationships with the Environment. Zoologisc her Anzeiger 240(3-4):525-533. Rebecchi, L and R Bertolani. 1994. Matura tive pattern of ovary and testis in eutardigrades of freshwater and terrestr ial habitats. Invertebrate Reproduction and Development 26:107-118. Rebecchi, L and R Bertolani. 1988. New cases of parthenogenesis and polyploidy in the genus Ramazzottius (Tardigrada, Hypsib iidae) and a hypothesis c oncerning their origin. Invertebrate Reproduction and Development 14:187-196. Rebecchi, L, T Abiera, and R Bertolan i. 1998. Chromosome C-banding and Ag-NOR pattern in tardigrades. Cy togent. Cell Genet. 81:27. Rebecchi, L and A Guidi. 1991. First SEM studies on tardigrade spematozoa. Invertebrate Reproduction and Development 19:151-156. Rebecchi, L and A Guidi. 1995. Spermatoz oon ultrastructure in two species of Amphibolus (Eutardigrada, Eohypsibiid ae). Acta Zoologica 76(2):171-176. Rebecchi, L and DR Nelson. 1998. Evaluation of a secondary sex characteristic in eutardigrades. Invertebra te Biology 117(3):194-198. Riggin, GT. 1962. Tardigrada of southwest Virgin ia: with the addition of a description of a new marine species from Flor ida. Virginia Agricultural Ex perimental Station, Technical Bulletin 152, 145 pp. Rho, HS, BH Min, and CY Chang. 1999. Taxono mic study of marine tardigrades from Korea. I. Genus Batillipes (Heterotardigrada:Batillipedidae). The Korean Journal of Systematic Zoology 15(1):107-118. Robinson, K, PJA Pugh, SJ McInnes, PJ Llewellyn, and SE Shackley. 1996. Cryopreparation of small or lightly attached biological specimen s. Microscopy and Analysis Nov:19.

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87 Appendices

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88 Appendix A Morphological Data Matrix Loricifera 1 1 1 1 1 1 1 1 1 1 1 1 1 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Kinoryncha 1 1 1 1 1 1 1 1 1 1 1 1 1 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Gastrotricha 0 0 0 1 1 1 1 1 1 1 1 0 0 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Macrobiotidae 1 1 1 1 1 1 0 0 0 1 1 1 1 1 0 0 2 2 1 2 2 2 1 1 1 2 3 4 2 2 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 2 1 0 0 Eohypsibiidae 1 1 1 1 1 1 0 0 0 1 1 1 1 1 0 0 ? 1 1 2 2 1 1 2 1 2 3 3 1 2 1 0 0 0 0 0 0 0 0 0 0 0 0 0 2 2 1 0 0 Calohypsibiidae 1 1 1 1 1 1 0 0 0 1 1 1 1 1 0 0 1 0 1 2 2 1 0 2 1 0 3 3 1 2 0 0 0 0 0 0 0 0 0 0 0 0 0 0 2 2 1 0 0 Necopinatidae 1 1 1 1 1 1 0 0 0 1 1 1 1 1 0 0 0 0 1 2 2 1 0 2 1 0 0 0 0 2 0 0 0 0 0 0 0 0 0 0 0 0 0 0 2 2 1 0 0 Microhypsibiidae 1 1 1 1 1 1 0 0 0 1 1 1 1 1 0 0 0 0 1 2 2 1 1 2 1 2 3 3 0 2 0 0 0 0 0 0 0 0 0 0 0 0 0 0 2 2 1 0 0 Hypsibiidae 1 1 1 1 1 1 0 0 0 1 1 1 1 1 0 0 2 2 1 2 2 1 1 2 1 0 3 3 2 2 2 0 0 0 0 0 0 0 0 0 0 0 0 0 2 2 1 0 0 Milnesiidae 1 1 1 1 1 1 0 0 0 1 1 1 1 1 1 0 1 1 1 2 0 0 0 2 1 0 2 2 0 2 0 0 0 0 0 0 0 0 0 0 0 0 0 0 1 2 1 1 0 Apodibius 1 1 1 1 1 1 0 0 0 1 1 1 1 1 0 0 0 0 1 2 0 0 1 2 1 0 0 0 0 2 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 2 1 0 0 Halechiniscidae 1 1 1 1 1 1 1 0 0 1 1 1 1 1 1 1 0 0 1 0 0 0 0 0 0 0 0 1 0 0 0 1 1 1 0 0 0 0 0 0 0 0 0 1 0 1 0 0 1 Stygarctidae 1 1 1 1 1 1 1 0 0 1 1 1 1 1 1 1 0 0 1 0 0 0 0 0 0 0 0 1 0 0 0 1 1 1 0 0 0 0 0 0 0 1 1 0 0 0 0 0 1 Renaudarctidae 1 1 1 1 1 1 1 0 0 1 1 1 1 1 1 1 0 0 1 0 0 0 0 0 0 0 0 1 0 0 0 1 1 1 1 1 0 1 1 1 0 1 1 ? 0 0 0 0 1 Coronarctidae 1 1 1 1 1 1 1 0 0 1 1 1 1 1 1 1 0 0 1 0 0 0 0 0 0 0 0 1 0 0 0 1 1 1 1 1 0 1 1 1 0 0 1 1 0 0 0 0 1 Batillipedidae 1 1 1 1 1 1 1 0 0 1 1 1 1 1 1 1 0 0 1 0 0 0 0 0 0 0 0 1 0 0 0 1 1 1 1 1 0 1 1 1 0 0 1 1 0 0 0 0 1 Echiniscoididae 1 1 1 1 1 1 1 0 0 1 1 1 1 1 0 1 0 0 1 0 0 0 0 0 0 0 0 1 0 0 0 1 0 0 0 0 0 0 0 0 0 1 1 0 0 2 0 0 1 Echiniscidae 1 1 1 1 1 1 1 0 0 1 1 1 1 1 0 1 0 0 1 0 0 0 0 0 0 0 0 1 0 0 0 1 0 1 1 1 1 1 1 1 2 1 1 ? 0 2 0 0 1 Oreellidae 1 1 1 1 1 1 1 0 0 1 1 1 1 1 0 1 0 0 1 0 0 0 0 0 0 0 0 1 0 0 0 1 0 0 0 0 0 0 0 0 0 0 0 ? 0 1 0 0 1

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89 Appendix B List of Morphological Characters (Unless otherwise stated, 0 = absent, 1 = present) (1) Molting by ecdysis. (2) Loss of locomotory cilia; Gastrotrichs ha ve locomotory cilia on their ventral side and they are covered by the cuticle. (3) Cuticle structure: trilaminate epicutic le, proteinaceous exocuticle and chitinous endocuticle; gastrotrichs do not have a chitinous endocuticle. (4) Parthenogenesis. (5) Circumpharyngeal nerve ring. (6) Complete gut. (7) Permanent genital pore separate from anus. (8) Adhesive glands. (9) Protonephridia. (10) Adult gut functional. (11) Triangular shaped pharynx. (12) Stylets: piercing rods lateral to the buccal tube that are anteriorly pointed and can be protruded out the mouth opening. (13) Formation of epicuticle: appears over th e tip of short microvilli in patches that laterally merge to form a continuous layer. In gastrotrichs the cutic le is brought to the surface by Golgi vesicles in the form of tiny plates. (14) Terminal mouth.

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90 (15) Cephalic papillae: short, rounded appenda ges occurring on the head, usually lateral to the mouth opening. (16) Cephalic appendages. (17) Peribuccal papillae: s hort, rounded appendages that surround the mouth opening. (18) Peribuccal lamellae: Short appendages surrounding the cuticular ring of the buccal opening (19) Buccal tube: anterior rigi d tube without spiral annulatio ns, extending from the mouth opening to the pharyngeal tube or the pharyx. (20) Buccal tube apophyses: cuticular thickeni ngs at the junction of the buccal tube and pharynx. (21) Pharyngeal tube: posterior flexible tube with spiral annulations, extending from the buccal tube into the pharynx. (22) Pharyngeal tube apophyses. (23) Ventral lamina: a small ventral support that extends from the mouth ring to approximately the middle of the buccal tube. (24) Stylet supports: structure that attaches the po sterior end of the stylet to the buccal tube. Stylet supports: 0 absent, 1 present, 2 either. (25) Macroplacoid: large, cuticular thickenings that occur in two or three transverse rows in the pharynx; Microplacoid: small, cuticul ar thickenings located posterior to the macroplacoids. Placoids: 0 absent, 1 mi croplacoids only, 2 macroplacoids only, 3 both micro and macroplacoids, (26) Septulum: cuticular thickenings at the base of the buccal tube. Septulum: 0 absent, 1 present, 2 either.

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91 (27) Claw Structure: 0 absent, 1 sing le, 2 double separated, 3 double connected. (28) Claw sequence, separa ted claws of the heterotard igrades were scored 1111 and Milnesiidae were scored 1122: 0 absent, 1 1111, 2 1122, 3 2121, 4 2112. (29) Transverse cuticular bar: 0 absent, 1 present, 2 either. Located at the base of the claws. (30) Accessory points: 0 absent, 1 present, 2 either. Located on the tips of the claws (31) Lunulae: 0 absent, 1 present, 2 either. (32) Lateral cirrus A: filame ntous appendages occurring at or near the junction of the head plate and the first segmental pl ate; associated with the clavae. (33) Median cirrus: filamentous appendages lo cated internally and/or ventrally to the cephalic papillae. (34) Cuticular armor: cuticular covering as in some heterotardigrades. (35) Dorsal segmental plates: unpaired plates located behind the head plate, and in the region of the first and second pair legs. (36) Head plate: the most anterior cuticula r plate, which bears th e cephalic appendages. (37) Median plate 1: the pl ate that is located between the first and second segmental plates. (38) Median plate 2: the plate that is lo cated between the second and third segmental plates. (39) Median plate 3: similar to median plates 1 and 2; loca ted between the third segmental plate and the end plat e or pseudosegmental plate. (40) Caudal plate: the most posterior cuticular plate.

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92 (41) Pseudosegmental plate: an unpaired plat e that is located imme diately posterior to median plate 3 and anterior to the end plate, as in Pseudechiniscus Pseudosegmental plate: 0 absent, 1 present, 2 either. (42) Peduncles. (43) Clavae: short, rounded appendages that occur at or near the junction of the head plate and the first segmental plat e; associated with cirri A. (44) Digitate legs. (45) Leg 4 morphology: Spines or papilla found on the 4th pair of legs: Leg 4 morphology: 0 absent, 1 spine, 2 papilla. (46) Eyespots: 0 absent, 1 present, 2 either. (47) Cloaca. (48) Sexual dimorphism of claws (49) Sexual dimorphism of the gonopore. (50) Pharyngeal stripes.

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About the Author Phillip Brent Nichols was born in Dyer sburg, Tennessee on the eleventh day of January nineteen hundred and seventy two. Th e only son of Bruce and Sandi Nichols of Moody, Alabama, he attended the Smith county school system and graduated from Smith County High School (Carthage, Tennessee) in May 1990. Upon completion of his M.S. degree in May 1999, Brent began working toward a Ph.D. at the University of South Florida in the lab of Dr. James R. Garey. While attending USF Brent instructed several course s in biology. He was an active member of the Graduate Assistants Union and was ac tive in re-establishing the Biology Graduate Student Organization and serv ed as its Vice-President fr om 2002-2003. He has published several papers from both his M.S. and Ph.D. degree work. Brent will enter the private work force upon graduation.


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Nichols, Phillip Brent.
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Tardigrade evolution and ecology
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by Phillip Brent Nichols.
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[Tampa, Fla] :
b University of South Florida,
2005.
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ABSTRACT: A character data set suitable for cladistic analysis of tardigrades at the family level was developed. The data matrix consisted of 50 morphological characters from 15 families of tardigrades and was analyzed by maximum parsimony. Kinorhynchs, loriciferans and gastrotrichs were used as outgroups. The results agree with the currently accepted hypothesis that Eutardigrada and Heterotardigrada are distinct monophyletic groups. Among the eutardigrades, Eoyhypsibiidae was found to be a sister group to Macrobiotidae + Hypsibiidae, while Milnesiidae was the basal eutardigrade family. The basal heterotardigrade family was found to be Oreellidae. Echiniscoideans grouped with some traditional Arthrotardigrada (Renaudarctidae, Coronarctidae + Batillipedidae) suggesting that the arthrotardigrades are not monophyletic. An 18S rRNA phylogenetic hypothesis was developed and supports the monophyly of Heterotardigrada and of Parachela versus Apochela within the Eutardigrada. Mapping of habitat preference suggest that terrestrial tardigrades are the ancestral state. Molecular analysis of a sediment sample with an unusually large population of tardigrades had a higher diversity when compared to manual sorting and counting.
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Dissertation (Ph.D.)--University of South Florida, 2005.
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Text (Electronic dissertation) in PDF format.
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Adviser: James R Garey.
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Ecdysozoa.
Meiofauna.
Phylogeny.
18s rrna.
Morphology.
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