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Graduate Theses and Dissertations
Graduate School
2005
Tardigrade evolution and ecology
Phillip Brent Nichols
University of South Florida
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Nichols, Phillip Brent, "Tardigrade evolution and ecology" (2005). Graduate Theses and Dissertations.
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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
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
Systematic Research
Dispersal
Ecological Importance
Significance
Objectives
1
5
9
10
11
13
Chapter Two – Specimen Acquisition and Processing
14
Chapter Three – Morphological Analysis of Tardigrade Phylogeny
Background
Material and Methods
Characters
Data Analysis
Results & Discussion
Morphological Phylogeny
Habitat Mapping
15
15
16
16
17
17
17
19
Chapter Four – Molecular Analysis of Tardigrade Phylogeny
Background
Material and Methods
Gene Selection
DNA Extraction
Polymerase Chain Reaction
Cloning
Sequencing
Alignments and Sequence Analysis
Results & Discussion
Molecular Phylogeny
Habitat Mapping
20
20
22
22
23
24
24
25
25
26
26
29
i
Chapter Five –Meiofaunal abundance, diversity and similarity in a uniquely rich
tardigrade community.
Background
Material and Methods
Sample Collection
DNA Sediment Extraction
Polymerase Chain Reaction
Cloning
Sequencing
Alignments and Sequence Analysis
Sediment Sorting and Specimen Identification
Data Analysis
Results & Discussion
Species Diversity
Community Similarity
31
31
34
34
34
35
36
36
37
37
37
38
48
48
Chapter Six– Summary
Morphological Analysis of Tardigrade Phylogeny
Molecular Analysis of Tardigrade Phylogeny
Meiofaunal Abundance, Diversity and Similarity in a Uniquely Rich
Tardigrade Community
51
51
51
References
53
Bibliography
62
Appendices
Appendix A. Morphological Data Matrix
Appendix B. List of Morphological Characters
86
88
89
About the Author
52
End Page
ii
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
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
Shannon (H’) and Simpson’s (C) diversity indices for molecular
OTU data and species groups.
48
Jaccard’s and Stander’s community similarity indices for molecular
OTU community compared to the manual count community.
49
Table 5.
Table 6.
Table 7.
iii
List of Figures
Figure 1.
Figure 2.
Figure 3.
Figure 4.
Figure 5.
Graphical representation of a typical tardigrade Redrawn from
Rammazzotti and Maucci (1983).
1
Typical tardigrade claws. A. Macrobiotus sp.; B. Milnesium sp.;
C. Echiniscus sp.; D. Batillipes sp. Redrawn from Rammazzotti
and Maucci (1983).
5
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
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
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
iv
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
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 8.
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
v
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
vi
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
Encystment
Description
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).
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 Ramazzotti and Maucci (1983).
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 (RenaudMornant, 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
Eutardigrada
Parachela
Calohypsibiidae
Eohypsibiidae
Hypsibiidae
Macrobiotidae
Microhypsibiidae
Necopinatidae
Milnesiidae
are dispersed
throughout three main
environments:
terrestrial, freshwater,
Apochela
and marine. Their
Heterotardigrada
Arthrotardigrada
distributions in each of
the habitats may be
correlated with
Echiniscoidea
physical
Batillipedidae
Coronarctidae
Halechiniscidae
Renaudarctidae
Stygarctidae
Echiniscidae
Echiniscoididae
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 (RodriguezRoda, 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.
13
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.
14
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)
17
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.
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.
19
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.
21
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).
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 intraphylum relationships
(Blaxter et al., 1998) and
that it has a long history
in determining interphylum relationships
Figure 7. This study used the 18S rRNA gene using primers
to amplify the portions of each gene as shown. See text for
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
were preserved in 50% glycerol and stored at –80 oC .
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
2002
1934
1918
2083
1872
X01723
X77786
U06478
X07801
U19182
Nelles et al., 1984
Chalwatzis et al., 1995
Campbell et al., 1995
Hendriks et al., 1988
Trapido-Rosenthal et al., 1994
1814
X53899
Rice 1990
1814
X80234
Winnepenninckx et al., 1995
1824
827
1783
1808
1735
1844
1777
1820
1771
1686
1432
1521
1384
1674
1726
AF056024
AY582118
AY582121
X81442
U32393
U49909
AY582120
AY582119
AY582122
AF056023
Present study
Present study
Present study
Present study
Present study
Garey et al., 1996
Jorgensen & Kristensen, 2004
Jorgensen & Kristensen, 2004
Giribet et al., 1996
Garey et al., 1996
Aguinaldo et al., 1997
Jorgensen & Kristensen, 2004
Jorgensen & Kristensen, 2004
Jorgensen & Kristensen, 2004
Garey et al., 1996
Present study
Present study
Present study
Present study
Present study
Arthropoda
Artemia salina
Meloe proscarabaeus
Okanagana utahensis
Tenebrio molitor
Panulirus argus
Mollusca
Placopecten magellanicus
Priapulida
Priapulus caudatus
Tardigrada
Echiniscus viridissimus
Halechiniscus remanei
Halobiotus stenostomus
Macrobiotus hufelandi
Macrobiotus tonolli
Milnesium tardigradum
Milnesium tardigradum
Pseudechiniscus islandicus
Ramazzottius oberhauseri
Thulinius stephaniae
Batillipes mirus
Hypsibius dujardini
Calohypsibius schusteri
Ramazzottius oberhauseri
Richtersius coronifer
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.
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. *Specimens were not identified to species in the original study, but have since been
verified.
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.
29
Figure 9. Tardigrade molecular topology from Fig. 8 with mapped habitat preference. Habitats:
Terrestrial (T ); Marine (M ); or Freshwater (F ).
30
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
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 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 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 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 ShannonWiener index (H’= -Σpi log pi) and Simpson’s index (C= 1-Σpi2), 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
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
S1140
Echiniscus
B1-10
Ballanus
S240
Plectus
66-18S4
Bugula
6718S4
Bugula
561864
Electra
40-18S4
Vaccinium
6018S4
Vaccinium
4418S4
Vaccinium
S381
Stenostomum
B1-9
Wilsonema
S2B7
Callinectes
S2B6
Aurila
S39
Caligus
39-18S4
Callinectes
S215
Daptonema
B1-8
Enoplus
B1-5
Chaetonotus
S341
Abyssothyris
S1B46
Brachionus
Total # of groups: 20
Common Name
Tardigrade
Barnacle
Nematode
Bryozoan
Bryozoan
Bryozoan
Plant
Plant
Plant
Flatworm
Nematode
Decapod
Ostracod
Copepod
Decapod
Nematode
Nematode
Gastrotrich
Brachiopod
Rotifer
Total # of individuals:
Frequency
27
1
7
1
1
28
1
1
12
3
1
1
6
5
1
8
1
8
6
8
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.
40
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.
44
45
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.
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
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
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
Tardigrade
Nematode
Barnacle
Polycheate
Flatworm
Rotifer
Diatoms*
Ostracod
Copepod
Gastrotrich
Molluscs
Crustacea
Brachiopod
Bryozoan*
Plant (Vaccinium)*
Total # groups: 15
Morphological
128
66
0
44
0
38
34
32
10
6
3
2
0
0
0
Total #: 363
OTUs
27
17
1
0
3
8
0
6
5
8
0
1
6
30
14
Total #: 126
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
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
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
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
98.6% similarity which is a function
of the number of species shared and
their relative distributions. The
Table 7. Jaccard’s and Stander’s community similarity
indices for molecular OTU community compared to the
manual count community.
Community Index
Jaccard’s
Stander’s
Similarity
0.583
0.986
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.
50
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.
52
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86
Appendices
87
Appendix A
Morphological Data Matrix
Loricifera
1111111111111100000000000000000000000000000000000
Kinoryncha
1111111111111100000000000000000000000000000000000
Gastrotricha
0001111111100100000000000000000000000000000000000
Macrobiotidae
1111110001111100221222111234221000000000000002100
Eohypsibiidae
1111110001111100?11221121233121000000000000022100
Calohypsibiidae
1111110001111100101221021033120000000000000022100
Necopinatidae
1111110001111100001221021000020000000000000022100
Microhypsibiidae
1111110001111100001221121233020000000000000022100
Hypsibiidae
1111110001111100221221121033222000000000000022100
Milnesiidae
1111110001111110111200021022020000000000000012110
Apodibius
1111110001111100001200121000020000000000000002100
Halechiniscidae
1111111001111111001000000001000111000000000101001
Stygarctidae
1111111001111111001000000001000111000000011000001
Renaudarctidae
1111111001111111001000000001000111110111011?00001
Coronarctidae
1111111001111111001000000001000111110111001100001
Batillipedidae
1111111001111111001000000001000111110111001100001
Echiniscoididae
1111111001111101001000000001000100000000011002001
Echiniscidae
1111111001111101001000000001000101111111211?02001
Oreellidae
1111111001111101001000000001000100000000000?01001
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.
92
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.

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