Tardigrade Evolution and Ecology Phillip Brent Nichols University of South Florida

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7-25-2005 Tardigrade Evolution And Ecology Phillip Brent Nichols University of South Florida

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This Dissertation is brought to you for free and open access by the Graduate School at Scholar Commons. It has been accepted for inclusion in Graduate Theses and Dissertations by an authorized administrator of Scholar Commons. For more information, please contact [email protected]. Tardigrade Evolution And Ecology

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

Dedication

For MaMa… Acknowledgements

There are so many people that I wish to thank and acknowledge for their love, support, and help over the past few years. First 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 more than one occasion. I will always value both the professional and personal friendship that we have. I would like to thank the members of my Graduate Supervisory Committee, 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, Heather Hamilton, and Stefie Depovic; have been involved with many aspects of this dissertation from collecting field samples to problem solving data collection. Other collaborators 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, University of Hawaii. I would like to extend my appreciation to the University of South Florida Graduate School and Biology

Department not only for financial support throughout my training but for all the “little things” that often go unnoticed. Lastly, I would like to thank my parents for providing me with the opportunities to pursue my own interests.

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

i 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 Identification 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 Morphological Characters 89

About the Author End Page

List of Tables

Table 1. Modes and causes of crytobiotisis in tardigrades. 4

Table 2*. Higher level Taxonomy of the Tardigrada. 9

Table 3. List of species, sequence length, origin, and sequence reference. 26

Table 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 Stander’s Similarity. 39

Table 5. Species groups and frequency of individuals per group. These data were used to calculate Shannon 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 community similarity indices for molecular OTU community compared to the manual count community. 49

iii

List of Figures

Figure 1. Graphical representation of a typical tardigrade Redrawn from Rammazzotti and Maucci (1983). 1

Figure 2. Typical tardigrade claws. A. Macrobiotus sp.; B. Milnesium sp.; C. Echiniscus sp.; D. Batillipes sp. Redrawn from Rammazzotti and Maucci (1983). 5

Figure 3. Representative head structures 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. Lateral 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. Macroplacoid; E. Microplacoid; F. Pharyngeal bulb Redrawn from Rammazzotti and 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 orders Parachela and Apochela along with the class Heterotardigrada; however, the orders within Heterotardigrada (Arthrotardigrada, Echiniscoidea) may not be monophyletic. Key to Characters: 1. Eutardigrada: No cephalic appendages; lack of dorsal plates; differentiated placoids; double 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. 18

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 phylogeny. 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. 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 claws but not digits (BARNES and HARRISON 1993). 6. Apochela: With cephalic papillae and with double 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 branches are joined (SCHUSTER et al. 1980). 8. Macrobiotus: Claw branches with sequence: secondary, primary, primary, secondary; buccal tube with ventral lamina and 10 peribuccal lamellae (SCHUSTER et al. 1980). 9. Thulinia + Hypsibius: Claw branches with sequence: secondary, 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). 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 numbers 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 sediment collection site on Dauphin Island, AL. 34

Figure 11. Rarefaction curve calculated for molecular OTUs and species groups. 39

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 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 phylogenetic tree was created based on the Kimura 2-parameter distance method and complete deletion of gaps. 45

Tardigrade Evolution and Ecology

Phillip Brent Nichols

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.

vii

Chapter One

Introduction

Tardigrades were first described by Goeze in 1773 and since then, his "Kleiner

Wasser Bärs" have been commonly referred to as “water-bears” because of their bear-like appearance. Spallanzani (1776) termed his similar organism "Il Tardigrado" (slow stepper) recognizing the unique lumbering gate exhibited by tardigrades. Tardigrades are found throughout the world in a variety of freshwater, marine, and terrestrial habitats and are considered cosmopolitan in their distribution. These bilaterally symmetric, hydrophilous micrometazoans are ventrally flattened and dorsally convex. A typical adult tardigrade (Figure 1) is 250-500 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 Figure 1. Graphical representation of a typical tardigrade. Redrawn from Ramazzotti and Maucci (1983). segments are present, each has a pair of ventrolateral legs terminating in claws or suction discs (Ramazzotti and

Maucci, 1983); generally the first three pairs are used for locomotion, the fourth for substrate attachment. Tardigrades have an outer cuticle that may be opaque, white, or 1 such colors as brown, green, pink, red, orange, 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 accomplished by piercing the cells with a pair of stylets and “sucking” out their contents, but in some cases whole organisms are ingested.

The typical digestive system is comprised of a foregut, midgut, and hindgut. The foregut includes a mouth, buccal tube, salivary glands, stylets, a sucking pharynx (with or without placoids), and an esophagus. The stylets 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 (Malpighian tubules) empty into the junction of the midgut and hindgut. The hindgut can either have a true cloaca (Eutardigrada) or an anus with a separate, preanal gonopore (Heterotardigrada).

Doyere proposed that all tardigrades were hermaphroditic based on what he identified as structures that were two testes and a seminal receptacle (Bertolani, 1979,

Nelson, 1982). This belief remained until two structures were identified as malpighian tubules and tardigrades were then considered to be gonochoristic (Bertolani, 1992;

Nelson 1982). However, Bertolani (1979) and Bertolani and Manicardi (1986) reported that hermaphroditism exists in some tardigrades.

2 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 females lay eggs outside the body or within the exuvium as they molt. The males then release spermatozoa into 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 gonochoristic tardigrades (amphimixis) can take place

between a female and a single male or several 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. Instead, mating habits have been

inferred from anatomical studies (Bertolani, 1992). Males and females are quite similar

but can be distinguished from one another by comparing the gonad and the gonoducts.

Females have one ovoduct whereas males have 2 vas deferens. Fertilization for terrestrial

species usually occurs inside the female’s body while in marine species fertilization is

external (Bertolani, 1990).

Parthenogenesis is the development of an egg where there has been no paternal

contribution of genes (Futuyma, 1986). This is the common mode of reproduction in unisexual tardigrades found in non-marine habitats (leaf litter, mosses, and freshwater)

(Nelson, 1982). Unisexual females are wide spread in tardigrades but recombination may be limited as they will have fewer possibilities for procreation. Parthenogenesis, while limiting genetic variability, may in fact facilitate proliferation in unisexual females

(Bertolani, 1987). Polyploidy, possessing more than one entire chromosomal compliment has often been associated with parthenogenesis and several species are known to have polyploid populations (Bertolani, 1982; Nelson, 1982). Also, cytotypes, 3 or populations with the same morphological 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 brain with two dorsolateral lobes connected by two circumpharyngeal commissures to a subpharyngeal ganglion, a pair of longitudinal nerve cords, and four ventral ganglia that are united by those nerve strands

(Dewel and Dewel, 1996).

The most fascinating feature of some tardigrades 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 barrel-shaped forming a tun. In this state,

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.

growth, reproduction, and metabolism are reduced or cease temporarily and resistance to environmental extremes is evident. This resistance allows the tardigrade to survive through cold and dry spells, ionizing radiation, heat, and pollution. Crowe (1975) identified five different types of cryptobiosis (Table 1).

4 Systematic Research

In 1928 and 1929 Marcus compiled the first reviews of tardigrade morphology,

physiology, embryology, and phylogenetic relationships; in 1936, he developed the first

basis for classification of the group (Kinchin, 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, Heterotardigrada, Eutardigrada, and

Mesotardigrada. Mesotardigrada was based on a single description of Thermozodium esakii from a hot spring near Nagasaki Japan. Type specimens were not preserved and the

hot spring where they were found was destroyed in an earthquake (Ramazzotti and

Figure 2. Typical tardigrade claws. A. Macrobiotus sp.; B. Milnesium sp.; C. Echiniscus sp.; D. Batillipes sp. Redrawn from Ramazzotti and Maucci (1983).

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 documented by Ramazotti and Maucci (1983) and

Kinchin (1994). Increased interest in tardigrade biology and systematics over the last 25

years is evidenced by numerous publications 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 5 morphological characters that include: claw size, shape, organization, and number

(Figure 2); organization of the bucco-pharyngeal apparatus; length of stylets; size, shape,

and number of placoids; cuticular patterns and 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 single 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 the 4th pair of legs. The Heterotardigrada are “armored” and

have a thick cuticle, dividing into plates, with species specific pore patterns. Two double

claws per leg, which may or may not be similar 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 characteristics are taxonomically significant.

6 The eutardigrades have a thin cuticle that is not divided into plates, but may be ornamented with spines, reticulations, 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 eutardigrades. The opening of the mouth is followed by the buccal tube which is supported by a ventral 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

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).

mouth by protractor and retractor muscles attached to the stylet supports. The buccal tube connects to the muscular pharyngeal bulb which in most tardigrades is lined with three rows of cuticular thickenings called placoids (macroplacoids and microplacoids). The placoids are posterior to the end of the buccal tube.

7 Usually gonochoristic, tardigrades have a single unpaired gonad but in many

cases the mode of reproduction is parthenogenic. 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 takes place inside. Some eutardigrade eggs are

freely laid and often ornamental. Both the 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 has often been debated (for reviews, see

Ramazzotti & Maucci, 1983; Kinchin, 1994), and morphology has primarily been used to

assess the relationships among some genera within a few families of tardigrades (Renaud-

Mornant, 1982; Kristensen, 1987; Pilato, 1989; Pollock, 1995; Bertolani & Biserov,

1996; Jorgensen, 2000; Guidetti & Bertolani, 2001). Tardigrades have been placed along an annelid-arthropod lineage, often closely 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 tardigrades, onychophorans, and arthropods form a monophyletic clade known as Panarthropoda (reviewed in Schmidt-Rhaesa et al., 1998).

Garey et al. (1999) found close agreement between molecular and morphology based phylogenies that included five species of tardigrades, suggesting 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,

8 the relationships among the families and among genera within the families have not been systematically evaluated. One exception is the family Echiniscidae for which several cladistic studies have been published (Kristensen, 1987; Jorgensen, 2000).

Dispersal Table 2*. Higher level Taxonomy of the Tardigrada Tardigrades Class Order Family are dispersed Eutardigrada Parachela Calohypsibiidae throughout three main Eohypsibiidae Hypsibiidae environments: Macrobiotidae Microhypsibiidae terrestrial, freshwater, Necopinatidae Apochela Milnesiidae and marine. Their Heterotardigrada Arthrotardigrada Batillipedidae distributions in each of Coronarctidae Halechiniscidae the habitats may be Renaudarctidae Stygarctidae correlated with Echiniscoidea Echiniscidae Echiniscoididae physical Oreellidae *(from Nelson, 2002) environmental factors.

The role of moisture, site orientation and altitude on the distribution of terrestrial tardigrade species has been widely investigated. Oxygen availability is the limiting factor in tardigrade distributions in all environments. It is because of oxygen availability that terrestrial tardigrades do not inhabit dense growing thick mosses and the reason that they are found in the top few centimeters of soil around the roots of trees, the soil must also have a degree of porosity to facilitate movement (Ramazzotti and Maucci, 1983).

Those in mosses and lichens, which have been categorized into types based on their

9 moisture content, need alternating 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 (Rodriguez-

Roda, 1951; Nelson, 1973, 1975; Ramazzotti and Maucci, 1983; Dastych, 1985, 1987,

1988; Beasley, 1988), with most suggesting that species richness increases 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 distributional patterns were not influenced by altitude (Kathman

and Cross, 1991).

Very little information exists as to the mode of dispersal of freshwater and marine tardigrades since very few undergo cryptobiosis. They are most likely distributed by way of rapid moving waters during periods of flooding, or by storm surge altering the coastal currents. Tardigrade zonation in littoral habitats, 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 tardigrades 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 consist of many diverse and abundant species of benthic invertebrates (including tardigrades). A typical food web often

10 consists of nematodes, tardigrades, bacteria, algae, rotifers, protozoa, mites, and collembolans (Kinchin 1987). One possible role of terrestrial tardigrades is in the colonizing of new habitats. Tardigrades may play an important role with the ability to move into a newly formed habitat quickly, establishing 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 and 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 food 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 areas 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) focused on community structure of marine species. Currently there are no models that can accurately measure their role in ecosystem functioning therefore, it would be impossible to assess their importance.

Significance

There currently is no phylogenetic hypothesis of tardigrade evolution that treats the group as a whole. Instead, the tardigrade literature consists of a plethora of species descriptions with very few systematic studies. A molecular framework for tardigrade

11 phylogeny would be very useful and would encourage a wider application of

morphological and molecular studies to tardigrade phylogeny. For example, a recent molecular study of nematode phylogeny (Blaxter et al., 1998) was pivotal in encouraging

new discussion and a reassessment of nematode 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 important 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.

12 Objectives

1. The present study was designed to provide an overall framework for tardigrade

evolution utilizing morphology and molecular based (18S rRNA) phylogenetic

hypotheses for tardigrades. Potential morphological characters that are informative at the

family level were assembled and a morphology 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 phylogenetic relationships among the

tardigrade families and to test the relationships among the families within each tardigrade order to determine if the families are truly monophyletic. Finally, it has been suggested that tardigrades evolved in a marine environment (May 1953) and that some echiniscids adapted to freshwater and then to a terrestrial 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 investigate the structure (frequency, diversity,

similarity) of a meiofaunal community that has an usually abundant tardigrade population

using both morphological species identification and molecular species identification.

Individuals were grouped to the lowest practical taxon and the abundance, diversity and similarity between the “morphological community” and “molecular community” was determined.

Chapter 2

Specimen Acquisition and Processing

Specimens were obtained from a network of tardigrade researchers and from field collections. Marine specimens were collected and rinsed with fresh water. The

freshwater rinse puts the tardigrades into osmotic shock and they release their grasp on

the sand grains. These samples were then rinsed through a set of nested sieves, which

have mesh openings from 1600mm to 40mm, with seawater to both collect the individual and stop the osmotic shock (Higgins and Thiel 1988). The specimens were drained into a screw-top jar and preserved by adding 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 agitated, removed, and squeezed to drain the

remaining water into the funnel. Samples were processed by sieving through nested

sieves. The specimens were then sorted, stored in screw-top jars, and preserved. The

cryptogams were re-dried and stored in previously 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.

Chapter 3

Morphological Analysis of Tardigrade Phylogeny

Background

Hennig (1979) stated that a classification should express the branching (cladistic)

relationships among species, regardless of their degree of similarity or difference.

Cladistic analysis is based on common ancestry 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 must determine the polarity of the character

states to show which are pleisiomorphic (ancestral) and which are apomorphic (derived).

Cladistic analyses utilize parsimony methods to construct a cladogram. This

method implies that the simplest answer is usually 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 represents the topology of a phylogenetic tree and can

be used to infer evolutionary history of the groups being studied. 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, parallelism and reversal are avoided. However, differing opinions

on character state definitions between researchers and the fact that fossil material may be

needed to evaluate a particular missing character can be a disadvantage.

15 The phylogenetic position of Tardigrada has 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 placed along an annelid-arthropod lineage, often closely associated with the onychophoran-arthropod 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 monophyletic clade known as Panarthropoda

(reviewed in Schmidt-Rhaesa et al., 1998).

The present study was designed to assemble 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 characters (Appendix B)

from current literature was scored for 15 tardigrade families (for review, see Schuster et

al., 1980; Binda & Kristensen, 1987; Kristensen, 1987; Bertolani & Rebecchi, 1993;

Schmidt-Rhaesa et al., 1998; Jorgensen, 2000; Guidetti & Bertolani, 2001). Characters

common to most families but with character states that distinguished families from one

another were chosen. Three outgroups; two within Ecdysozoa (Loricifer & Kinorhyncha)

16 and one outside Ecdysozoa (Gastrotricha) were included in the analysis. Ten multi-state

characters were present.

Data Analysis

The character data matrix was constructed using Nexus Data Editor (NDE)

version 0.5.0 (http://taxonomy.zoology.gla.ac.uk/rod/NDE/nde.html). Phylogenetic

Analysis Using Parsimony (PAUP*) version 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 sister groups as evidenced by a high bootstrap

value of 98%. Among the eutardigrades, Eohypsibiidae was a sister group to

Macrobiotidae + Hypsibiidae, though the bootstrap support around 60% was minimal.

Necopinatidae appears to be basal among the Parachela, which forms a monophyletic

sister group to the Apochela and the most basal eutardigrade family, Milnesiidae (Figure

5).

Among the heterotardigrade families, bootstrap values indicate that all branches

were moderately or highly supported (Figure 6). Kristensen & Higgins (1984)

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

18 heterotardigrade family. The current analysis (Figure 5) suggests instead that

Halechiniscidae is more basal than either Stygarctidae or Renaudarctidae and that

Oreelidae is the most basal heterotardigrade. This placement supports earlier suggestions by Thulin (1928) and Marcus (1929) that Echiniscidae is derived from arthrotardigrades

and that Oreellidae should be considered the most primitive heterotardgrade. The

cephalic appendages that define heterotardigrades are present in oreellids, but the dorsal

plates, characteristic of other heterotardigrades, are completely absent. The morphology

of the various dorsal plates appears to provide important characters in heterotardigrade

phylogeny. If the absence of dorsal plates represents a secondary loss within the

oreellids, then the basal position of Oreellidae in this study may be an artifact.

The orders Arthrotardigrada and Echiniscoidea have been placed as sister groups

based on shared derived characters (Eibye-Jacobsen, 2001). The current results (Figure

5) suggest that these orders are not sister groups, and it appears 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 environments twice, to freshwater environments at

least three times, and to terrestrial environments twice. The placement of the terrestrial

Oreellidae as the basal heterotardigrade suggests that tardigrades were ancestrally

terrestrial. However, the placement of Oreellidae in the character tree may be an artifact

caused by the secondary loss of dorsal plates.

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 clade that includes arthropods.

There appears to be a consensus that arthropods, 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 arthropod outgroup (Regier & Shultz, 2001). Other molecular studies have placed tardigrades within a group of molting animals that includes arthropods, priapulids, nematodes and kinorhynchs (Aguinaldo et al., 1997; Zrzavy et al.,

1998). Garey et al. (1999) found close agreement between molecular and morphology based phylogenies that included six species of tardigrades, suggesting that the characters for tardigrade morphological studies are appropriate. This study also suggested that

Heterotardigrades, the marine and terrestrial armored species, are the most basal group

with the greatest number of plesiomorphic characters.

Garey et al. (1996) showed that tardigrades are closely allied to arthropods and

that tardigrades, arthropods and priapulids form a clade. Since priapulids had long been

20 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 aschelminths while other characters such as body

muscle specialization, lack of a closed circulatory 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 together in studies of rotifer phylogeny, tardigrades, protostomes, urochordates, 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) demonstrated that trees produced by the analysis of 18S rRNA gene sequences of tardigrades are in close agreement to trees based on morphological characters (Fig. 6). This suggests that molecular 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.

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 considered and rejected for this study because of the small sample size and the relative difficulty in obtaining both DNA

22 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 intra-

phylum relationships

(Blaxter et al., 1998) and

that it has a long history

in determining inter- Figure 7. This study used the 18S rRNA gene using primers to amplify the portions of each gene as shown. See text for phylum relationships details.

among metazoans (Field et al., 1988).

DNA Extraction

Tardigrades were isolated onto individual glass slides and air dried. The

tardigrade cuticle was then mechanically macerated using dental probes previously

cleaned with DNA-Away (Molecular BioProducts, Inc., San Diego, CA) and rinsed in

deionized water. A 10 µl solution containing 5mg/ml of Proteinase K was added to the

macerated cuticle and the sample was treated 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 shown to improve the efficiency of cloning

in PCR products (Crowe et al., 1991). The extraction mixture was then transferred to a

200 µl 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 µl of the supernatant was then used as

template DNA.

23 Polymerase Chain Reaction

PCR amplification of template DNA was carried out using primers specific for the

18S rRNA gene (18S4 5'-GCTTGTCTCAAAGATTAAGCC-3' and 18S5 5'-

ACCATACTCCCCCCGGAACC-3'). PCR reactions were performed in 200 µl tubes

using a Biometra TRIO thermocycler (WhatmanBiometra, Göttingen, Germany). The

reaction cycle consisted of an initial 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 µM final concentration of each primer, 0.25 mM final concentration for each of dATP, dCTP, dTTP, and dGTP, 10 µl of template DNA and 1 unit of EnzyPlus

2000 Taq polymerase (Enzypol, Denver, CO) in a final volume of 50 µl.

Cloning

PCR product of 18S rDNA amplified from some of the samples was cloned using

the TOPO TA cloning kit for sequencing (Invitrogen Corp., San Diego, CA) following

the manufacturer’s instructions. Transformed cells were plated and incubated overnight at

370C on Luria-Bertani (LB) agar containing 100µg/mL ampicillin and 50µg/mL X-gal

(5-Bromo-4-chloro-3-indolyl β-D- galactoside). 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 Isolation Kit and then quantified using 0.9%

agarose gel electrophoresis. All colonies grown overnight for isolation of plasmid DNA

o were preserved in 50% glycerol and stored at –80 C .

24 Sequencing

Template DNA (cloned and direct PCR product) was cycle sequenced using the

QuickStart sequencing kit (BeckmanCoulter, La Jolla, CA). The reaction mix contained

between 50 and 100ng of template, 2 µl sequencing reaction mix (SRM), 1µl 3.2 µM

sequencing primer, and water to bring the total reaction volume to 10 µl. All products

(clones and PCR) for analysis were sequenced using the 18S4c and 18S2c primer.

Sequencing reactions were amplified in the Biometra TRIO thermocycler. A preheat step

consisting of only water and template 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 seconds and an extension step

at 60 oC for one minute. Finally, a 60 oC extension step for 7 minutes was performed and

a final hold at 4 oC. The cycle sequencing product was purified following the

manufacturer’s instructions and analyzed using a CEQ 8000 Genetic Analysis system

(BeckmanCoulter, La Jolla, CA).

Alignments and Sequence Analysis

Sequences were visually checked and corrected for ambiguous bases (N’s). Data

sets from individual tardigrades were aligned according to a secondary structure model

(Neefs et al. 1993) using DCSE (De Rijk and De Wachter 1993). Phylogenetic analysis

of alignments was performed using MEGA version 2.1 (Kumar, et al. 2001) to produce

neighbor-joining trees and Phylogenetic Analysis Using Parsimony (PAUP*) version 4.6

(Swofford, 1998) was utilized for Maximum Parsimony analysis. A variety of arthropod

25 species were used as outgroups. Trees produced 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 length, 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

26 The NJ and MP trees produced from 18S rRNA gene sequences (Figure 8) were congruent with one another and support the morphological character analysis in indicating that Eutardigrada and Heterotardigrada are each monophyletic as evidenced by

bootstrap values of 89 and 84, respectively.

27 The 18S rRNA gene sequences from Calohypsibius schusteri, Hypsibius

dujardini, Richtersius coronifer, and Ramazzotius oberhouseri adds 4 additional genera

to the previously published eutardigrade dataset. 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 statistical support for relevant nodes within

Eutardigrada in the molecular results (82% - 100% bootstrap values) than in the

morphological results (54% - 64% bootstrap values). This analysis agrees with the

findings of Jorgensen and Kristensen (2004) that the eutardigrades are monophyletic and

it is consistent with the morphological analysis in Figure 5. However, the taxon sampling

among the Macrobiotidae family is not exhaustive and some questions about the

monophyly of this line still 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 level Murrayidae. Nearly 40% of the Parachela

genera are currently within the family Macrobiotidae and this is clearly an area that needs

further study in order to determine an accurate phylogeny.

28 Within heterotardigrades, the monophyly of Arthrotardigrada and Echiniscoidea

is well supported with a bootstrap value of 84%. There is more statistical support for

relevant nodes in the molecular results (90% - 100% bootstrap values) than in the

morphological tree (60% - 64% bootstrap values). The 18S rRNA gene sequence from

Batillipes mirus and Halechiniscus remanae (Arthrotardigrada) adds an additional

heterotardigrade order and two new families (Batillipedidae, Halechiniscidae) to the

previously published molecular data set. The addition of Pseudechiniscus islandicus

adds an additional genus within the Echiniscidae family to the dataset. The molecular data suggests that heterotardigrades and eutardigrades are both monophyletic.

Habitat Mapping

As seen in the morphological dataset, mapping of habitat preferences onto the gene tree (Figure 9A) also suggests that tardigrades have adapted to marine environments

twice, to freshwater environments once, and to terrestrial environments twice. The

placement of B. mirus and H. remanaei as the basal heterotardigrade suggest that

tardigrades were ancestrally marine. By mapping the habitats preferences on the tree we

can trace the number of evolutionary events that occur. If terrestrial tardigrades are

considered to be ancestral state then there are four events that occur (Figure 9B) while if

we assume that the marine tardigrades are the ancestral tardigrades then there are five events that occur (Figure 9C). This suggests 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.

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

Chapter 5

Meiofaunal Abundance, diversity and Similarity in a

Uniquely Rich Tardigrade Community.

Background

Meiofauna are distributed worldwide and include representatives of 22 of the

metazoan phyla (Coull, 1999, 1988). They are found in terrestrial, freshwater and marine

habitats and are classified by their small size, generally 50 - 500 µm. Ecologically,

meiofauna function as food sources for higher trophic 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 sediment characteristics (particle size) and

bacterial production (Higgins, 1988; Vanreusel et al., 1995; Schratzberger et al., 2000) and are often reported in terms of abundance, distribution and diversity (eg. Hummon,

1987; Trett et al., 2000, Battigelli & Berch, 2004).

Meiofaunal assemblages in marine habitats 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 depth and salinity gradients.

Typically, the highest abundances are found in intertidal zones and decrease as the depth

31 increases. Salinity, food availability, sediment grain size, and tidal exposure can affect dispersal (Findlay, 1981) which is often patchy with densities changing over a distance of just a few centimeters (Schratzberger et al., 2000).

Surveys on marine tardigrade distributions are primarily descriptions of new species and taxonomical reviews (De Zio Grimaldi & D’Addabbo, 2001). Studies investigating the ecology of marine tardigrades most often do not report information on their abundance and ecology within the context of the meiofaunal community. A few exceptions are from ecological studies from the Mediterranean Sea (for review see, De

Zio Grimaldi & D’Addabbo, 2001), psammolittoral tardigrades from North Carolina

(Lindgren, 1971), interstitial tardigrades 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 meiofaunal assemblages and used them as a measure of the health of the environment (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 identifications and the need for broad taxonomic expertise (Warwick, 1988), an area of study that has declined in recent years.

Traditional sorting techniques have involved 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 sediment extracts may provide a more cost

32 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 environmental samples (Stephen et al., 1996) and those that

have been cultured represent only a fraction of the estimated species (Wintzingerode et

al., 1997). Techniques developed to extract DNA from sediment samples have allowed microbial ecologists to analyze the species composition of unculturable microbes

(Kennedy and Gewin, 1997). The techniques developed 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 similarity 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 investigate the diversity using rRNA sequence data.

They were able to determine groups and identify 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 numbers 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

33 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 meiofaunal DNA from marine sediment collected on

Dauphin Island, Alabama. Sediment samples were 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 site 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.

34 The samples were then homogenized using pre-cleaned Teflon tipped pestles and

centrifuged to pellet the sediment. The supernatant was pipetted to a 1.5 mL centrifuge

tube and an equal volume of phenol (pH 7.9) was 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 transferred to a new 1.5mL tube and the previous steps were repeated: one more time using phenol (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 and 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 rpm for 15 minutes, washed

with 70% ethanol and suspended in 100 µl of deionized water.

Polymerase Chain Reaction

PCR amplification of template DNA was 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 µl tubes

using a Biometra TRIO thermocycler (WhatmanBiometra, Göttingen, Germany). The

reaction cycle consisted of an initial 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 µM final concentration of each primer, 0.25 mM final concentration for

35 each of dATP, dCTP, dTTP, and dGTP, 10 µl of template DNA and 1 unit of EnzyPlus

2000 Taq polymerase (Enzypol, Denver, CO) in a final volume of 50 µl.

Cloning

PCR product of was cloned using the TOPO TA cloning kit for sequencing

(Invitrogen Corp., San Diego, CA) following the manufacturer’s instructions.

Transformed cells were plated and incubated overnight at 37 oC on Luria-Bertani (LB)

agar containing 100µg/mL ampicillin and 50µg/mL X-gal (5-Bromo-4-chloro-3-indolyl

β-D- galactoside). 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 Isolation Kit and then quantified using 0.9% agarose gel electrophoresis. All colonies grown overnight for isolation of plasmid DNA were

o preserved in 50% glycerol and stored at –80 C .

Sequencing

Template DNA (cloned and direct PCR product) was cycle sequenced using the

QuickStart sequencing kit (BeckmanCoulter, La Jolla, CA). The reaction mix contained

between 50 and 100ng of template, 2 µl sequencing reaction mix (SRM), 1µl 3.2 µM

sequencing primer, and water to bring the total reaction volume to 10 µl. All products

(clones and PCR) for analysis were sequenced using the 18S4 and 18S5 primer.

Sequencing reactions were amplified in the Biometra TRIO thermocycler. A preheat step

consisting of only water and template 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,

36 denature step for 15 seconds, a 50 oC annealing step for 30 seconds and an extension step at 60 oC for one minute. Finally, a 60 oC extension step for 7 minutes was performed and a final hold at 4 oC. The cycle sequencing product was purified following the manufacturer’s instructions and analyzed using a CEQ 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 analysis of the alignment was performed using MEGA version 2.1 (Kumar et al., 2001) to produce neighbor- joining 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 differences 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 specimen identification

Meiofaunal samples preserved in 95% ethanol with Rose Bengal stain were manually sorted and counted using a Meiji dissecting scope. Type of species present and frequency of individuals were keyed to the most 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. This dataset was aligned with ClustalX. A

37 phylogenetic tree utilizing the neighbor-joining method and the Kimura 2-parameter

distance model was generated. The OTUs were 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 diversity was calculated using the Shannon-

2 Wiener index (H’= -Σpi log pi) and Simpson’s index (C= 1-Σpi ), where pi is the number

of individuals of a species divided by the total number of individuals. Community similarity was calculated using 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 respectively; and proportional similarity.

The two communities compared in this study are defined as 1) the OTUs from the

sequence analysis and 2) the manual 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 yielded 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 reference data set and unique sequences

representing each of the OTUs identified twelve groups (Table 4). Hamilton (2003,

unpublished thesis) showed that specific sequences of meiofaunal data could be identified

from a reference alignment. The frequencies of individual sequences from each of the 12

groups identified here are reported in Table 4.

38 Table 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 (Table 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

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

41 Figure 12 (Continued)

42 Figure 12 (Conitinued)

43 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.

45 Figure 13 (Continued)

46 Species groups from manual sorting were determined and the frequencies of individuals present are reported in Table 5. Ten groups were found. Tardigrades were the dominant species Table 5. Morphological and sequence group (OTUs) comprising nearly 35% of the frequency of individuals. These data, with noted omissions (*), were used to calculate Shannon diversity, population. Nematodes Simpson’s Diversity, Jaccard’s Similarity and Stander’s Similarity. (18%), polychaetes (12%), Species Morphological OTUs Tardigrade 128 27 rotifers (10%), diatoms (9%) Nematode 66 17 Barnacle 0 1 Polycheate 44 0 and ostracods (8%) were the Flatworm 0 3 Rotifer 38 8 other main taxa identified. Diatoms* 34 0 Ostracod 32 6 There is a striking contrast in Copepod 10 5 Gastrotrich 6 8 Molluscs 3 0 the groups represented when Crustacea 2 1 Brachiopod 0 6 compared to the molecular Bryozoan* 0 30 Plant (Vaccinium)* 0 14 Total # groups: 15 Total #: 363 Total #: 126 data set. Bryozoans, which were absent in the manual counts, and tardigrades 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 Table 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

47 fragments of bryozoan and pollen from the asterids. 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 series of populations that are equally abundant

will have high diversity indices, while those 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 sensitive 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 morphological groups (Table 6). The number of

Table 6. Shannon (Hmax) and Simpson’s (C) diversity species present in OTUs (n=10) and indices for molecular OTU data and species groups Index OTUs Species groups morphological groups (n=9) differs by

Shannon (H’) 0.837 0.733 a count of 1. The distribution of Simpson’s (C) 0.231 0.176 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

48 communities. Two methods of calculating community similarity are Jaccard’s coefficient

and Stander’s coefficient. Jaccard’s coefficient 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 Table 7. Jaccard’s and Stander’s community similarity indices for molecular OTU community compared to the 98.6% similarity which is a function manual count community. Community Index Similarity of the number of species shared and Jaccard’s 0.583 Stander’s 0.986 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 discriminating between the meiofaunal groups

(OTUs). The results indicate that rarefaction curves can reach saturation and that groups

can be identified from small sample sizes. These results can then be used to compare

49 communities utilizing species diversity, community similarity and other comparative

analyses. The capability to identify specific groups utilizing gene sequences may lead to

more detailed surveys of meiofaunal communities by lessening the burden on the

researcher attempting to identify individuals through traditional techniques.

Tardigrades from the family Batillipedidae are 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 meiofauna 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 presence of a sand bar, which is unique when

compared to a typical salt marsh. The interaction 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 area 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 investigations looking for tardigrades in intertidal salt

marshes.

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 Parachela is a monophyletic

sistergroup to Apochela within the eutardigrades while the order Parachela is a

monophyletic sister group to the Apochela. The phylogeny also suggests that the

heterotardigrade orders Arthrotardigrada and Echiniscoidea 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. However, 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 supported and the molecular data showed high support for the relationship between the families. The monophyly of the Macrobiotidae family remains in question. The molecular phylogeny, in contrast to the morphological

51 phylogeny, showed that the order Arthrotardigrada and Echiniscodea are monophyletic

groups. Mapping of the habitat preferences suggests that terrestrial tardigrades are the ancestral state. However, more extensive taxon sampling will be required to test this hypothesis.

Meiofaunal Abundance, diversity and Similarity in a Uniquely Rich Tardigrade Community

The molecular and phylogenetic methods used here to assess a community of marine tardigrades were successful in discriminating between the meiofaunal groups

(OTUs) identifiying 17 OTUs in contrast to the 12 morphological groups 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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Appendices

87 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

88 Appendix B

List of Morphological Characters

(Unless otherwise stated, 0 = absent, 1 = present) (1) Molting by ecdysis.

(2) Loss of locomotory cilia; Gastrotrichs have locomotory cilia on their ventral side and they are covered by the cuticle.

(3) Cuticle structure: trilaminate epicuticle, 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 the tip of short microvilli in patches that laterally merge to form a continuous layer. In gastrotrichs the cuticle is brought to the surface by Golgi vesicles in the form of tiny plates.

(14) Terminal mouth.

89 (15) Cephalic papillae: short, rounded appendages occurring on the head, usually lateral to the mouth opening.

(16) Cephalic appendages.

(17) Peribuccal papillae: short, rounded appendages that surround the mouth opening.

(18) Peribuccal lamellae: Short appendages surrounding the cuticular ring of the buccal opening

(19) Buccal tube: anterior rigid tube without spiral annulations, extending from the mouth opening to the pharyngeal tube or the pharyx.

(20) Buccal tube apophyses: cuticular thickenings 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 posterior 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, cuticular thickenings located posterior to the macroplacoids. Placoids: 0 absent, 1 microplacoids 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.

90 (27) Claw Structure: 0 absent, 1 single, 2 double separated, 3 double connected.

(28) Claw sequence, separated claws of the heterotardigrades 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: filamentous appendages occurring at or near the junction of the

head plate and the first segmental plate; associated with the clavae.

(33) Median cirrus: filamentous appendages located 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 cuticular plate, which bears the cephalic appendages.

(37) Median plate 1: the plate that is located between the first and second segmental plates.

(38) Median plate 2: the plate that is located between the second and third segmental plates.

(39) Median plate 3: similar to median plates 1 and 2; located between the third segmental plate and the end plate or pseudosegmental plate.

(40) Caudal plate: the most posterior cuticular plate.

91 (41) Pseudosegmental plate: an unpaired plate that is located immediately 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 plate; 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.

About the Author

Phillip Brent Nichols was born in Dyersburg, Tennessee on the eleventh day of

January nineteen hundred and seventy two. The 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 courses in biology. He was an active member of the Graduate Assistants Union and was active in re-establishing the Biology Graduate

Student Organization and served as its Vice-President from 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.