Kenneth Halberg

Osmoregulation | With Focus on Fluid
and Solute Dynamics in Tardigrada
PhD Dissertation
KENNETH A. HALBERG
© Kenneth A. Halberg

FACULTY OF SCIENCE
UNIVERSITY OF COPENHAGEN
Osmoregulation│With Focus on Fluid
and Solute Dynamics in Tardigrada
PhD Dissertation
Kenneth A. Halberg
UNIVERSITAS
HAFNIENSIS
2012
Dissertation submitted Monday the 14th of May 2012.
Supervisor: Associate Professor Nadja Møbjerg, PhD.

Dissertation presented at University of Copenhagen to be publicly examined (provided acceptance in its
current form) in Auditorium 1, August Krogh Building, Universitetsparken 13, Thursday, June 28, 2012
at 14:00 for the degree of Doctor of Philosophy. The examination will be conducted in English.
Abstract
Halberg, K. A. 2012. Osmoregulation │With Focus on Fluid and Solute Dynamics in
Tardigrada.
Osmoregulation is the regulated control of water and solute composition in body fluid
compartments. On one hand, the internal composition must be kept within optimal
conditions for metabolic processes in the face of external perturbation. On the other
hand, the nature of the living state demands a continuous traffic of compounds in and
out of the organism. These demands appear to be in fundamental contradiction
however cells and animals achieve so-called “steady-state” by means of an array of
transport proteins, which provide a stringent control on the exchange of water and
solutes across body surfaces. The distinct mechanisms of solute transport have been
studied in most animal groups, but there are still large gaps in our understanding of
how animals cope with osmotic stress. In the present thesis, osmoregulatory
phenomena were studied in vertebrate and invertebrate organism alike, with the main
focus being on fluid and solute dynamics in Tardigrada. For example, the inorganic
ion composition of several species was investigated, which revealed that tardigrades
contain roughly similar relative contributions of inorganic ions to total osmotic
concentration, when compared to closely related animal groups. Moreover, it was
inferred that cryptobiotic tardigrades (species able to enter a state of latent life)
contain a large fraction of organic osmolytes. The mechanisms of organic anion
transport in a marine species of tardigrade was investigated pharmacologically, and
compared to that of insects. These data showed that organic anion transport is
localized to the midgut epithelium and that the transport is both active and transporter
mediated with a pharmacological profile similar to that of insects. Tardigrades survive
in a variety of osmotic environments (semi-terrestrial, limnic and marine habitats),
why the ability to volume and osmoregulate was examined. These studies demonstrated
an ability to regulate total body volume during both hypo- and hyperosmotic
conditions, and that the ability to hyper-regulate could be a general theme among
members of eutardigrades. Thus, the work presented herein, have contributed to
establishing tardigrades as an important experimental group in which central
physiological questions may be answered, including aspects of osmotic and ionic
regulation.
Keywords: osmoregulation, volume regulation, organic anion transport, hyper-regulate,
inorganic ions, organic osmolytes, tardigrade, insect,
Kenneth A. Halberg, The August Krogh Centre, Department of Biology,
Universitetsparken 13, DK-2100 Copenhagen Ø, Denmark
© Kenneth A. Halberg 2012

“Beautiful is what we see,
More beautiful is what we perceive,
Most beautiful is what we do not understand”
- Niels Stensen

List of Papers
This thesis is based on the following papers and manuscripts, which are referred to
in the text by their Roman numerals.
I. Halberg, K. A., Larsen, K. W., Jørgensen, A., Ramløv, H. & Møbjerg,
N. Cryptobiotic tardigrades contain large fraction of unidentified
organic solutes: A comparative study on inorganic ion composition in
Tardigrada.
II.
Halberg, K. A., & Møbjerg, N. (2012). First evidence of epithelial
transport in tardigrades: A comparative investigation of organic anion
transport. Journal of Experimental Biology, 215: 497-507.
III. Møbjerg, N. M., Halberg, K. A., Jørgensen, A. Persson., D., Bjørn M,
Ramløv H & Kristensen R. M. (2011). Survival in extreme
environments – on current knowledge of adaptations in tardigrades.
Acta Physiologica, 202: 409-420.
IV. Haugen, B.M., Halberg, K.A., Jespersen, Å., Prehn, L.R. & Møbjerg,
N. (2010). Functional characterization of the vertebrate primary ureter:
Structure and ion transport mechanisms of the pronephric duct of
axolotl larvae (Amphibia). BMC developmental Biology, 10: 56.
V. Halberg, K. A., Persson, D., Ramløv, H., Westh, P., Kristensen, R. M.
& Møbjerg, N. (2009). Cyclomorphosis in Tardigrada: Adaption to
environmental constraints. Journal of Experimental Biology, 212:
2803-2811.
Additionally, the following papers and manuscripts were prepared during the
course of my PhD studies, but are not included in the thesis:
VI.
VII.
Halberg K. A., Jørgensen, A. and Møbjerg, N. (in prep.). Surviving
without water: Tun formation in tardigrades is an active process
mediated by the musculature
Halberg K. A., Persson, D., Jørgensen, A. Kristensen, R. M. and
Møbjerg, N. (submitted). Population dynamics of a marine tardigrade:
Temperature limits geographic distribution of Halobiotus crispae.
Marine Biological Research

VIII. Persson, D., Halberg K. A., Jørgensen A., Møbjerg N. & Kristensen
R. M. (in review). Neuroanatomy of Halobiotus crispae (Eutardigrada:
Hypsibiidae): Tardigrade brain structure suggests inclusion into
Panarthropoda. Journal of Morphology.
IX. Persson, D., Halberg K. A., Jørgensen A., Ricci C., Møbjerg N. &
Kristensen R. M. (2010). Extreme stress tolerance in tardigrades:
Surviving space conditions in low earth orbit. Journal of Zoological
Systematics and Evolutionary Research, 49: 90-97.
X. Halberg, K. A., Persson D., Møbjerg N., Wanninger A. & Kristensen
R. M. (2009). Myoanatomy of the Marine Tardigrade Halobiotus
crispae (Eutardigrada: Hypsibiidae). Journal of Morphology, 270:
996-1013.
Lastly, following paper provides important background knowledge for the work
presented herein:
XI. Møbjerg, N., A. Jørgensen, J. Eibye-Jacobsen, K. A. Halberg, D.
Persson & R. M. Kristensen (2007). New Records on cyclomorphosis
in the marine eutardigrade Halobiotus crispae (Eutardigrada:
Hypsibiidae). Journal of Limnology, 66 (suppl. 1): 132-140.

Reprint and publication is made with permission from the respective copyright
holders.
Paper II, V © The Company of Biologists.
Paper IV, is copyright of the authors.
Paper III © Wiley-Blackwell
Statement of authorship
Paper I: KAH was deeply involved in study design. KAH participated in
extracting animals and ion chromatography. KAH performed nanoliter osmometry.
KAH performed the data analysis, prepared the figures, participated in discussions
and interpretation of the data, and drafted the manuscript.
Paper II: KAH performed the major part of the experimental work and data
analysis. He participated in planning of experiments, data interpretation, prepared
the figures and drafted the manuscript.
Paper III: KAH performed cell counts and provided images of tardigrades. He
helped draft parts of the manuscript.
Paper IV: KAH participated in immunostaining experiments, performed CLSM
and prepared the 3D images. KAH participated in discussions and interpretation of
the data.
Paper V: KAH participated in planning of experiments, sampling, staging,
scanning electron microscopy, DSC experiments, experiments on cold hardiness
and osmotic stress tolerance, volume measurements, hemolymph sample
collections, and nanoliter osmometry. He furthermore participated in discussions
and interpretation of data, prepared the figures and drafted the manuscript.
Front cover: Scanning elecron micrographs of the tardigrades Rictersius coronifer
(top left), Halobiotus crispae (middle right), and Echiniscus testudo (middle
bottom).

Contents
Preface.......................................................................................................................... 9
Introduction ................................................................................................................ 11
Maintaining a stable internal environment .......................................................... 11
Osmoregulators and osmoconformers ................................................................. 12
Osmoregulatory organs........................................................................................ 12
Filtration-Reabsorption systems ................................................................ 13
Secretion-Reabsorption systems ................................................................ 14
Phylum Tardigrada..................................................................................................... 17
General morphology ............................................................................................ 18
Classification ....................................................................................................... 19
Ecology ................................................................................................................ 21
Fluid and solute dynamics – an overview .................................................................. 23
Inorganic ion composition ................................................................................... 23
Organic anion transport ....................................................................................... 25
Volume and osmoregulation................................................................................ 26
Conclusions and future perspectives .......................................................................... 27
Dansk sammenfatning ................................................................................................ 29
Acknowledgements .................................................................................................... 30
References .................................................................................................................. 32

Preface
The primary aim of this thesis was to address several aspects of the fluid and
solute dynamics in tardigrades, and hereby provide new insight into the general
stress biology of these enigmatic creatures. This was done by adopting an
integrative approach, i.e. applying advanced methods in biology, analytic
biochemistry and physical chemistry, which offered functional data from different
disciplines to be obtained. The experimental work was carried out mainly at The
August Krogh Centre, University of Copenhagen; however, additional
experimental work was performed at the Danish Natural History Museum,
University of Copenhagen and at the Department of Nature, Systems and Models,
Roskilde University. Overall this thesis has contributed to converting tardigrades,
from an almost exclusive taxonomic phenomenon into an established and
important experimental group, in which central physiological questions can be
investigated.
This PhD-thesis comprises a short introduction to osmotic and ionic
regulation in Metazoa, accompanied by a brief review on the general morphology,
classification and ecology of tardigrades. Moreover, an overview of the results
presented as well as conclusions and future perspectives are presented. Five papers
and manuscripts form the basis of this thesis, of which four are published in peer
review journals (Papers II, III, IV and V), and one is prepared for submission
(Paper I). I am the first author of three (Paper I, II and V) and second author on
two (Papers III and IV) of these papers. Five additional papers and manuscripts
were prepared during the course of my PhD studies, but are not included in the
thesis. This work was funded by the 2008 Faculty of Science, University of
Copenhagen Freja-Programme.
Copenhagen, the 14 th of May 2012
Kenneth A. Halberg
9

Introduction
Maintaining a stable internal environment
“Life is as a thing of macromolecular cohesion in salty water” (Gilles & Delpire,
1997). Albeit a crude statement, it frames the fact that the ability to control salt
and water balance is a fundamental prerequisite for both cellular and animal life.
Indeed, the internal environment must usually be kept within relatively narrow
limits, as substantial deviations in cell composition are incompatible with the
optimal function of macromolecules (lipids, proteins, RNA), and may ultimately
modify the rate and extent of cellular reactions (Zhao, 2005). The overall
mechanism by which animals conserve a proper osmotic balance between cells,
extracellular fluid and the environment is termed osmoregulation.
The basis for osmoregulation lies in the strict control of the ionic
composition and the osmotic pressure of the intra- and extracellular compartments
through the regulated accumulation and loss of inorganic ions and organic
compounds (Dawson & Liu, 2009). This regulation is achieved through the
coordinated activity of an array of transporter proteins (both energy-consuming
and passive), which collectively maintain the steady-state condition of cells and
animals (Essig, 1968). During steady-state conditions, compositions of the intraand
extracellular compartments are maintained in a non-equilibrium state (Dawson
& Liu, 2009). This uneven distribution of solutes is important for keeping an
optimal milieu for metabolic processes (Zhao, 2005). Accordingly, the
extracellular fraction of the body fluids of animals are typically high in Na + and Cl -
, and relatively low in the other major ions (e.g. K + , Ca 2+ and Mg 2+ ), while the
intracellular environment of most organisms is low in Na + but high in K + , PO 4
3-
and proteins (e.g. Dawson & Lui, 2009; Paper I). As such, the plasma membrane
of cells must maintain ionic, but not osmotic, differences, while specialized
excretory organs – e.g. antennal glands of crustaceans, Malpighian tubules of
insects and tardigrades, rectal glands of sharks and rays, gills and intestine of
teleost fishes, salt glands of birds and reptiles, the kidneys of vertebrates etc. –
often maintain both ionic and osmotic differences between animals and their
environments (Riegel, 1970; Peaker, 1971; Paper IV, Beyenbach & Piermarini,
2011; Reilly et al., 2011; Whittamore, 2012). In general, mechanisms that allowed
organisms to respond and adapt to an osmotic challenge over the course of
11

evolution, has been fundamental to the invasion of new osmotic hostile habitats,
and such ecological divergence in turn is an important mechanism for the
speciation process (Schluter, 2009). Accordingly, if the ability to osmoregulate
had not evolved, life on the planet would look quiet different from how we know
it.
Osmoregulators and osmoconformers
Generally, animals are divided into two broad categories in terms of their
responses to osmotic stress: osmoregulators, which maintain an internal osmolarity
different from that of the external environment, and osmoconformers, which
conform to the external medium in which they are immersed (Fig. 1). Most
vertebrates are strict osmoregulators (e.g. Paper IV), as they maintain ionic and
osmotic balance within narrow limits; although hagfish, a basal group of
vertebrates, are a notable exception (Sardella et al., 2009). Conversely, marine
invertebrates are typically categorized as osmoconformers, as many of them
appear to be in osmotic balance with sea water over a range of salinities (Fig. 1).
However, there are numerous obvious exceptions including the marine tardigrade
Halobiotus crispae as well as several members of Crustacea (e.g. Sarver et al.,
1994; Normant et al., 2005; Paper V). In fact, the terms ‘strict osmoregulator’ and
‘strict osmoconformer’ must be used with caution, as typical osmoregulators are
forced to conform during the most extreme conditions (e.g. Dowd et al., 2010),
whereas animals otherwise described as osmoconformers actually maintain slight
differences between the internal and external environment (e.g. van Weel, 1957).
Although achieved through different mechanisms, both osmoregulating and
osmoconforming animals may tolerate wide ranges of external salinities, thus
termed euryhaline species, while animals intolerant of large changes are called
stenohaline species.
Osmoregulatory organs
Osmoregulatory organs are specialized organs involved in maintaining ionic and
osmotic homeostasis in the face of osmotic perturbation, as well as in excreting
endobiotic and exobiotic waste products (Dawson & Lui, 2009; Papers II, IV).
The specific organs mediating these processes may vary between different groups
of animals (see above); however, the molecular basis and specific mechanism of
solute and water transport often show a highly convergent/homologous pattern
across different types of epithelia (e.g. Paper II). In general, two types of systems
12

Fig. 1 Examples of osmotic performance of representative species from various groups
exposed to sea/brackish water, expressed as the relation between internal (extracellular) and
external (habitat) osmolality. These data show that osmoconformaty (∆osm=0) is present in
invertebrate and vertebrate species alike, albeit strong hypo-regulators and the ability to
produce a hyperosmotic urine appears restricted to vertebrates. It should be emphasized that
the selected species not necessarily represent the osmotic performance of the entire group, as
large difference may exist between even closely related species. For original data see:
Robertson, 1949 a ; van Weel, 1957 e ; Dice, 1968 d ; Liggins and Grigg, 1985 j ; Diehl, 1986 b ;
Normant et al., 2005 f ; Sardella et al., 2009 g ; Paper V c ;Reilly et al., 2011 h ; Whittamore,
2012 i ).
have evolved by which the initial process of urine formation takes place i) the
filtration-reabsorption type and ii) the secretion-reabsorption type.
Filtration-Reabsorption systems
The kidneys of vertebrates (fish, amphibians, reptiles, birds and mammals), and
the functional analogs of crusteans and molluscs, maintain extracellular fluid
homeostasis by producing urine through the filtration of plasma (ultrafiltration),
which is subsequently modified by selective reabsorption and secretion of ions,
organic molecules and water (Anderson, 1960; Schmidt-Nielsen, 1963; Riegel,
13

1970; Møbjerg et al., 2004; Paper IV; Whittamore et al., 2012). In vertebrates,
three temporally and spatially different kidney generations, the pronephroi,
mesonephroi and metanephroi, successively maintain fluid and electrolyte
homeostasis during morphogenesis, with the pronephroi constituting the functional
kidneys of fish and amphibian larvae (Paper IV). The functional unit of the
vertebrate kidney is the nephron, which is composed of a filtration unit and a renal
tubule (Anderson, 1960; Møbjerg et al., 2004; Paper IV). The filtration process
that takes place in the filtration unit (the glomerulus and Bowman’s capsule) is
‘passive’ i.e. entirely driven by the hydrostatic pressure generated by the heart,
whereas the reabsorption and secretion processes take place over specialized
epithelia of the renal tubule (Schmidt-Nielsen, 1963). The primary membrane
transporter for energizing vertebrate tissue is the Na/K-ATPase (e.g. Paper IV).
Accordingly, vertebrate kidneys may produce both dilute, iso-osmotic and
concentrated urine relative to the body fluids (Paper IV; Whittamore et al., 2012),
which has been a dominant factor in allowing vertebrates to penetrate into all types
of habitats on Earth. Filtration-reabsorption systems are capable of processing
large volumes of fluids, but are energetically costly, as any substance (e.g.
glucose) that has been filtered remains in the urine unless subsequently reabsorbed
(Schmidt-Nielsen, 1963). The advantage of such a system; however, is that new
potentially toxic compounds are eliminated without the need for developing
distinct secretory pathways for each new compound, which may be necessary for
the secretion-reabsorption type system (Paper II).
Secretion-Reabsorption systems
The Malpighian tubules of insects, and possibly tardigrades (Møbjerg & Dahl,
1996; Papers II, III, V), are the functional analogs of the vertebrate kidney, but
constitute a secretion-reabsorption system that produces urine in a fundamentally
different way than the filtration-reabsorption-systems (Beyenbach & Piermarini,
2011). In the absence of blood vessels (i.e. a closed circulatory system), the
hemolymph of insects is circulated at pressures insufficient for filtration, and the
Malpighian tubules thus form the (primary) urine entirely by secretion (Beyenbach
& Piermarini, 2011). The formation of the primary urine is generally initiated in
the distal segments (blind-ended tip) of the Malpighian tubule, and is essentially
iso-osmotic (consisting mainly of KCl and NaCl) to the hemolymph (Williams &
Beyenbach, 1983). The subsequent reabsorption of water, ions and metabolites in
proportions that maintain extracellular homeostasis (a process analogous to that of
vertebrates) takes place in downstream structures i.e. proximal tubule, hindgut and
rectum (O’Donnell & Maddrell, 1995; Coast, 2007). The final urine composition is
14

adjusted in the rectum (Coast, 2007) and may be either strongly hypo- or
hyperosmotic depending on the species and its physiological status (Maddrell &
Phillips, 1975; Reynolds & Bellward, 1989). In contrast to vertebrate epithelia, the
V-type H + -ATPase is considered ubiquitous in energizing insect epithelia
(Beyenbach & Piermarini, 2011); although the Na/K-ATPase is still expressed and
functionally relevant for tubular function (Torrie et al., 2004; Paper II). In fact,
energized by the H + -ATPase, some of the highest fluid secretion rates per unit area
membrane from any tissue have been reported from hematophagous insects (e.g.
Aedes aegypti, Rhodnius prolixus) after a blood meal (Williams & Beyenbach,
1983; Maddrell & Phillips, 1975). In addition to playing a key role in
osmoregulation, new properties of the Malpighian tubules of insects have emerged
in recent years, which suggest that Malpighian tubules are involved in such diverse
functions as renal detoxification, metabolism of toxins and immune system
responses (Dow & Davies, 2006; Paper II).
As holds for insects, ultrastructural studies on the Malpighian tubules of
tardigrades indicate that they function as secretion-reabsorption systems involved
in fluid and solute transport (Weglarska, 1987; Møbjerg and Dahl, 1996; Peltzer et
al., 2007). They are positioned at the transition zone between the midgut and
rectum of eutardigrades (Fig. 2), and the positional conformity between insects
and eutardigrades has been used as a strong argument for their homology (Greven,
1982; Møbjerg and Dahl, 1996). However, at present no functional data exist
relating the Malpighian tubules of tardigrades to an osmoregulatory role.
Accordingly, functional studies on the fluid and solute dynamics of tardigrades are
greatly needed (Papers I, II, III, V), and due to the close affinity to the
euarthropod complex (Aguinaldo et al., 1997), would be highly useful in
understanding and reconstructing the evolution of osmoregulation in Insects and
other arthropods.
15

Fig. 2 Structure and organization of the Malpighian tubules of the marine eutardigrade
Halobiotus crispae. dm, dorsal Malpighian tubule; dp, distal part; is, initial segment; mg,
midgut; mv, microvilli,; nu, nucleus; pp, proximal part; re, rectum; From: Møbjerg & Dahl,
1996.
16

Phylum Tardigrada
Tardigrades, also known as water bears, are among the smallest multi-cellular
animals on the planet (0.1-1.2 mm). They were discovered in 1773 by the German
pastor J. A. E. Goeze, who described them as “kleiner Wasserbär”, or little water
bear, due to their strong resemblance to a tiny bear (Ramazzotti & Maucci 1982).
Not long after the current name “Tardigrada” was given by the Italian naturalist
Spallanzani in 1776 (Lat. tardus – slow, grado – walker). In response to
unfavorable environmental conditions, many species of tardigrades have the ability
to enter the ametabolic state of suspended animation, also known as cryptobiosis,
in which the organism is neither dead nor alive (Møbjerg et al., 2011; Fig. 3). The
animal can remain in this state for as much as 20 years (Jørgensen et al., 2007), yet
once external conditions again become favorable, the tardigrade resumes activity
unaffected. This incredible ability is shared with selected species of nematodes,
rotifers and arthropods (Glasheen & Hand, 1988; Crowe & Maddin, 1974; Ricci et
al., 2003). In 1962, Tardigrada was recognized as a phylum by Ramazzotti in Il
Phylum Tardigrada. Presently, there are more than 1000 described species
(Guidetti & Bertolani 2005; Degma & Guidetti, 2007; Degma et al., 2012);
however, it has been estimated that several thousand species remain undescribed
(Paper III).
Tardigrada is included in the invertebrate superclade Ecdysozoa
(Aguinaldo et al. 1997); however, their precise phylogenetic position is still being
debated. Both molecular and morphological investigations produce conflicting
conclusions, and it is currently unclear whether the group is more closely related to
the nematodes and nematomorphs or to arthropods and onychophorans (Aguinaldo
et al. 1997; Dunn et al. 2008; Zantke et al., 2008; Edgecombe 2010; Rota-Stabelli
et al., 2010; Cambell et al., 2011). Regardless, this group is closely related to one
of the two most species-rich and economically important groups Nematoda or
Arthropoda, and thus maintains a central position in relation to the two major
invertebrate model organisms, i.e. Caenorhabditis elegans Maupas, 1900 and
Drosophila (Sophophora) melanogaster Meigen, 1830 (Gabriel et al. 2007;
Goldstein and Blaxter 2002).
17

Fig. 3 Scanning electron micrographs showing the external morphology of Richtersius
coronifer (Eutardigrada) A. lateral view of the active, hydrated state B. Dorsal view of the
dehydrated, cryptobiotic state C. Ventral view of the dehydrated, cryptobiotic state. The
pictures illustrate the morphological changes associated with entry into an ametabolic state
(i.e. cryptobiosis), which include the retraction of head and limbs into the body cavity, and the
formation of a compact shape – the tun. From: Paper VI.
General morphology
Tardigrades are bilaterally symmetric micrometazoans with a body divided into
five separate body segments, i.e. a cephalic segment, containing a mouth, eyespots
18

and sensory organs (papillae cephalica or cirri and clavae), and four trunk
segments (Nelson, 2002; Fig. 3A). The first three trunk segments each bear a pair
of lateroventrally directed legs, while the terminal trunk segment bears a pair of
posterioventrally directed legs (Figs. 3, 4). The legs typically terminate in 4 to 13
claws or suction discs (Nelson, 2002). Tardigrades are ventrally flattened with a
convex dorsal side, and are covered by a segmented cutinous cuticle, which is
periodically shed during molting – formation of the new cuticle is maintained by a
single layer of epidermal cells (Nelson, 2002). The digestive system consists of a
foregut, midgut and a hindgut with a pair of stylets and stylet glands flanking the
buccal tube. Three glands (the Malpighian tubules) are positioned at the transition
zone between the midgut and hindgut in eutardigrades (Weglarska, 1987; Møbjerg
and Dahl, 1996; Peltzer et al., 2007). Tardigrades posses a hemocoel-type of fluidfilled
body cavity, i.e. an open circulatory system as seen in arthropods and
nematodes, which likely functions in circulation and respiration. The somatic
musculature is composed of structurally independent muscle fibers, which can be
divided into a dorsal, ventral, dorsoventral, and a lateral musculature in addition to
a distinct leg musculature (Schmidt-Rhaesa & Kulessa, 2007; Fig. 4). Moreover,
the buccopharyngeal muscles, intestinal muscles and cloacal muscles comprise the
animal’s visceral musculature. Whereas cross striation of the somatic musculature
is especially pronounced in Arthrotardigrada, the somatic muscles of Eutardigrada
are described as an intermediate between smooth and obliquely striated (Walz,
1974). The nervous system of tardigrades consists of an (at least) three lobed brain
(Fig. 5) and a ventral nerve cord with four fused paired ganglia that shows a clear
segmental organization.
Classification
Originally based on morphological characters, tardigrades are divided into two
main evolutionary lines, represented by the extant lineages Eutardigrada and
Heterotardigrada. The validity of a third class, Mesotardigrada, is currently
considered dubious (Ramazzotti and Maucci, 1983).
Heterotardigrada consists of the orders Arthrotardigrada and Echiniscoidea
with arthrotardigrades possessing the most plesiomorphic characters.
Arthrotardigrada consists exclusively of marine species (Renaud-Mornant 1982;
Jørgensen et al. 2010), and are morphologically the most diverse group. They are
present in all oceans from intertidal zones to abyssal depths, and inhabit various
types of sediment. In contrast, the Echiniscoidea comprises both limno-terrestrial,
limnic as well as marine species with the majority of the described species
belonging to the family Echiniscidae. Taxonomically, the main characters
19

Fig. 4 Tardigrade musculature as revealed by fluorescently coupled phalloidin in combination
with confocal laser scanning microscopy three-dimensional reconstruction A. Lateral view of
Halobiotus crispae (Eutardigrada), Paper VIII B. Ventral view of Echiniscoides sigismundi
(Heterotardigrada), unpublished data.
separating the two groups from Eutardigrada include a separate gonopore, a closed
three-lobed anus as well as well-developed cephalic-, trunk-, and leg appendages
(Guidetti and Bertolani, 2005).
The eutardigrades are divided into the two orders; Apochela and Parachela.
Both orders contain mainly limno-terrestrial species, albeit with a few exceptions –
20

Fig. 5 Conceptual drawing based on immunofluorescent and ultrastructural data, showing an
interpretation of the brain structure of Halobiotus crispae. A. Lateral view B. Frontal view.
clg, claw gland; co, connective; dc, dorsal commissure; ey, eye; g0, sub-pharyngeal ganglion;
gI, first ventral trunk ganglion; ic, inner connective; il, inner lobe; lgg, leg ganglion; mg,
median ganglion; mo, mouth opening; oc, outer connective; ol, outer lobe; pc, papilla
cephalica; pb, pharyngeal bulb; st, stylet; t, temporalia; vll, ventrolateral lobe. From: Paper
VIII.
most notably the marine genus Halobiotus (see Paper III). In general,
eutardigrades are cylindrically shaped with a more or less distinct segmentation,
and exhibit a relatively uniform morphology (Fig. 3A). The key characters of
eutardigrades include a cloaca (combined gonopore and anus), the presence of
Malpighian tubules and a strong reduction of cephalic sensory structures (Guidetti
and Bertolani, 2005). Structures such as the bucco-pharyngeal apparatus and claw
shape are important taxonomic characters within Eutardigrada.
Ecology
Tardigrades occupy a range of moisture regimes and often constitute a major
component of meiofaunal communities in terrestrial, limnic and marine
ecosystems throughout the globe. However, they are distinctly aquatic organisms,
requiring a film of water to be active. Tardigrades are predominantly egg-laying,
with both sexual and parthenogenetic modes of reproduction described (Bertolani,
2001). Molting occurs continuously throughout their lifecycle, which may be
between 3-30 months (Nelson, 2002). Populations of tardigrades have been
studied in a variety of habitats; including mosses, lichens, leaf litter and soil, and
21

Fig. 6 Population dynamics of the marine tardigrade Halobiotus crispae showing a unimodal
pattern of maximal frequency. Graphic representation of sampling data (2006-2012)
comparing the temporal pattern in abundance of H. crispae to abiotic parameters
(temperature, (─ ─); salinity, (- - -); pH, (- ─ -), and the seasonal appearance of the different
cyclomorphic stages (shown on top), from the locality of Vellerup Vig, Isefjord, Denmark.
Light grey area is the period in which exuvia containing eggs were found, and thus indicates
the period of sexual reproduction. From: Paper VII.
the life history and population dynamics have received some attention (Martinez,
1975; Morgan 1977; Guidetti et al. 1999; Uhía and Briones 2002; Jönson 2003;
Suzuki 2003). Tardigrade population dynamics may show both unimodal and
bimodal patterns of annual variation (Martinez, 1975; Morgan, 1977; Fig. 6),
albeit the specific pattern appears to be both species and habitat specific. Factors
such as temperature, moisture and food availability have been suggested to be
correlated with population density (Hallas & Yeates, 1972; Morgan, 1977).
However, other factors including competition, predation and parasitism may play a
role in controlling population density and diversity (Nelson, 2002). Tardigrades
possess an amazing reproductive capacity, as indicated by the large changes in
animal density on short temporal scales (Morgan, 1977).
22

Fluid and solute dynamics – an overview
Inorganic ion composition
Knowledge of the composition as well as concentrations of dissolved particles in
internal fluids of an organism, and how these change during various exposures, is
fundamental to the understanding of its basic physiology. However, practically
nothing is known about these aspects in tardigrades, which has been a major
obstacle to the study of the fluid and solute dynamics in these animals. In Paper I,
the inorganic ion content of five different species (Echiniscus testudo, Milnesium
tardigradum, Richtersius coronifer, Macrobiotus cf. hufelandi and Halobiotus
crispae) covering both a large phylogenetic and habitat spectrum was analyzed
(Fig. 7A). These data demonstrated that Na + and Cl - are the principle inorganic
ions in tardigrade fluids, albeit substantial concentrations of K + , NH + 4 , Ca 2+ ,
Mg 2+ , F - 2-
3-
, SO 4 and PO 4 were also detected. Moreover, tardigrades appear to
contain roughly similar relative contributions of the respective inorganic ions to
total osmotic concentration, when compared to selected species of arthropods,
nematodes and onychophorans (Hobson et al., 1952; Sutcliffe, 1962; Campiglia,
1975; Wilder et al., 1998; Normant et al., 2005), albeit a large relative contribution
of calcium appears characteristic of tardigrade fluids. Inorganic ions destabilize
macromolecules and affect the rate and extent of metabolic reactions at high
concentrations, which invariably leads to impairment of cellular function (Zhao,
2005; Yancey, 2005). However, apparently R. coronifer does not exclude
inorganic ions during dehydration (Fig. 7B), which suggests a concomitant
accumulation of organic solutes. Moreover, as evidenced by the differences
between the calculated osmotic concentrations of the known ions and the
measured total osmolarity in the different species (Fig. 7A), it was inferred that
cryptobiotic tardigrades (in steady-state) contain a large fraction of unidentified
organic osmolytes. Organic osmolytes can be divided into a few major categories
(sugars, polyols, amino acids and various derivatives), and several of these groups
possess known protective functions in relation to osmotic stress (Hincha and
Hagemann, 2004; Yancey, 2005). Accordingly, the future detection and analysis of
such compounds are likely to provide new insight into the biochemistry and
physiology of superior tardigrade adaptations.
23

Fig. 7 Ionic contributions to total osmotic concentration A. Concentrations (mM) of the
respective cations and anions measured in each investigated species, as well as the
corresponding total osmotic concentration (mOsm/kg). The blank area represents the osmotic
deficit (OD). B. Concentrations (mg/l) of the respective cations and anions measured in
hydrated, active specimens compared to dehydrated cryptobiotic animals of Richtersius
coronifer. Data are expressed as mean ± s.d. From: Paper I.
24

Organic anion transport
The ability to excrete endogenous
waste products as well as environmental
toxins is an essential step to
avoid these compounds reach toxic
levels and to keep metabolic reactions
within optimal conditions. One of the
better known systems involved in such
an excretion is the organic anion
transport system, the function of
which has been studied in several
vertebrate (fish, amphibians, reptiles,
birds and mammals) and invertebrate
(nematodes and insects) model
organisms (George et al., 1999;
Dantzler, 2002; Dow and Davies,
2006). In Paper II, these data were
expanded as the sites, characteristics
and pharmacological profile of the
transepithelial transport of chlorophenol
red (CPR), a prototypical
Fig. 8 Tentative cellular model for the
transepithelial transport of organic anions in
tardigrades. From: Paper II.
substrate of the classic organic anion secretion pathway, was investigated in the
tardigrade Halobiotus crispae Kristensen, 1982 and compared to corresponding
data from the desert locust Schistocerca gregaria Forskål, 1775. This was done by
introducing a new method for quantifying non-fluorescent dyes. Our study
revealed i) that tardigrades posses an organic anion transport system, ii) that it was
localized to the midgut epithelium, and iii) that organic anion secretion was both
active and transporter mediated, with possible members of the SLC21/SLCO
transport families mediating the basolateral entry step in tardigrade midgut cells.
Transport by insect Malpighian tubules showed a similar pharmacological profile,
but higher concentrations of CPR were achieved. Based on the observed transport
characteristics in the presence and absence of transport inhibitors, a tentative
cellular model for the transepithelial transport of CPR in tardigrades was
suggested (Fig. 8). Specifically, a large lumen positive potential generated by the
H + -ATPase could provide the driving force for accumulation of anions in the
lumen, although the exact coupling between electrochemical gradients generated
by the pumps and transport of ions is unknown. This study was the first to provide
evidence for epithelial transport in tardigrades.
25

Volume and osmoregulation
Tardigrades survive in a variety of osmotic environments and must protect the
internal tissues from the vagaries and extremes of the external environment. The
secondary marine species Halobiotus crispae colonizes habitats characterized by
especially large fluctuations in external salinity, why the ability to respond to an
osmotic challenge was investigated in this species (Paper V). Animals were
exposed to both hypo- and hyperosmotic media and the subsequent changes in
total body volume and internal osmotic pressure were recorded. These data
revealed that H. crispae is able to regulate total body volume back to control
values when immersed in hypotonic solutions, yet was unable to do so in
concentrated media. Instead a new steady-state was achieved significantly below
control conditions. Animal activity was only markedly affected in very dilute
media, suggesting a possible effect on neuro-muscular function at low salt
concentrations. Conversely, when analyzing the concomitant changes in
hemolymph concentrations, it appeared that H. crispae is a euryhaline
osmoconformer, in which the hemolymph osmotic pressure is largely governed by
the external environment. However, expressing the hemolymph osmolality
measured during exposure to dilute as well as concentrated media as a function of
external salinity revealed that H. crispae is in fact a strong hyper-regulator (Fig.
9A). Expanding these studies to include the limno-terrestrial species Richtersius
coronifer Richters, 1903 (Paper III) showed that this is could be a general feature
of all eutardigrades (Fig. 9B), which was further supported by data presented in
Paper I. The ability to hyper-regulate indicates the excretion of dilute urine. The
three glands positioned at the transition zone between the midgut and rectum of
eutardigrades, are generally considered to have an excretory function. This
assumption is based on the probable homology of the Malpighian tubules of
eutardigrades and insects (Greven, 1982; Møbjerg and Dahl, 1996), and on several
ultrastructural studies that shows that the epithelium is likely involved in fluid and
solute transport (Weglarska, 1987; Møbjerg and Dahl, 1996; Peltzer et al., 2007).
However, data in support of an osmoregulatory function of both the rectum
(Dewel and Dewel, 1979) and midgut (Paper II) also exists, which emphasizes the
need for functional studies on these organs at both the molecular and cellular level.
26

Fig. 9 Osmotic performance of A. Halobiotus crispae and B. Richtersius coronifer during
exposure to media of varying osmotic strength. From: Paper III.
27

Conclusions and future perspectives
This dissertation has provided new insight into the fluid and solute dynamics of
metazoans, particularly relating to one of the most enigmatic groups on the planet
– the tardigrades. Using a multi-disciplinary approach, crucial information was
provided on both organs and systems of several species, representing vertebrates,
arthropods and tardigrades, and general patterns in especially tardigrade
physiology have emerged (e.g. Paper III). By comparing our data on tardigrades
to several evolutionary related groups, including nematodes, onychophorans and
arthropods, basic physiological principles have been discovered (e.g. Paper II),
which emphasizes the importance of comparative physiology.
In this respect, future work on tardigrade physiology could encompass an
extension of the work presented herein. Additional species from different habitats
and evolutionary lineages should be investigated, in order to further explore the
diversity as well as common trends in tardigrade biology. In particular, studies on
marine cryptobionts (e.g. Echiniscoides sigismundi) and additional noncryptobiotic
species would help clarify whether the osmotic deficits observed in
cryptobiotic animals (Fig. 5; Paper I) in fact are related to cryptobiotic ability or
alternatively to habitat preference. Regardless, large scale analyses and
characterization of the organic solutes of cryptobiotic tardigrades should be
performed, which surely would provide an enhanced resolution of several aspects
relating to tardigrade stress responses. Other studies could include a
characterization of the volume and osmoregulatory capacity of heterotardigrade
species. Our results on eutardigrades show a capacity to hyper-regulate over a
broad range of external salinities (Fig. 7); however, do heterotardigrades without
Malpighian tubules posses the same ability True limnic species (e.g. Bertolanius
nebulosus) should also be investigated. In addition to the suggested whole animal
experiments, studies on the molecular and cellular level are needed to fully
understand how fluid and electrolyte homeostasis is achieved in these animals. For
example, electrophysiological investigations using single cell glass microelectrode
impalements on dissected native tissue would help characterize the cellular
transporters involved in e.g. urine formation. Collectively, these studies would be
important to elucidate how tardigrades functionally have solved colonizing every
major type of habitat on the planet, and perhaps be useful in reconstructing how
osmoregulation has evolved in metazoans.
28

Dansk sammenfatning
Osmoregulering er kontrollen af kropsvæskernes sammensætning af vand og
opløste stoffer. Foruden at opfylde de optimale betingelser for metaboliske
processer under indflydelse af eksterne påvirkninger, skal denne
sammensætning samtidig imødekomme den kontinuerlige transport af stoffer
ind og ud af den levende organisme. Disse krav forekommer umiddelbart
fundamentalt uforenelige, men celler og dyr opnår et såkaldt ”steady-state” ved
hjælp af et spektrum af transportproteiner, der udøver streng kontrol over
udvekslingen af vand og opløste stoffer over forskellige kropsoverflader. De
forskellige transportmekanismer ansvarlige for denne kontrol er blevet
undersøgt i de fleste dyregrupper, men vores viden om hvorledes dyr håndterer
osmotisk stress, er stadig ufuldstændig. I dette arbejde er osmoregulatoriske
fænomener blevet undersøgt i såvel vertebrater som invertebrater, omend der
hovedsageligt er blevet fokuseret på dynamikkerne af vand og opløste stoffer i
Tardigrada. Eksempelvis blev sammensætningen af uorganiske ioner undersøgt
i flere forskellige arter af bjørnedyr, hvilket afslørede, at de relative bidrag af
uorganiske ioner til den totale osmotiske koncentration overordnet set er ens i
bjørnedyr og nært beslægtede dyregrupper. Desuden blev det udledt, at
kryptobiotiske bjørnedyr (arter, der er i stand til indtræde i et stadie af latent
liv) indeholder en stor andel af organiske osmolytter. Mekanismerne for
aniontransport blev undersøgt farmakologisk i en marin art af bjørnedyr, og
sammenlignet med de tilsvarende mekanismer i insekter. I bjørnedyret blev den
organiske aniontransport lokaliseret til epitelet i midttarmen, og viste sig at
være en aktiv transport, med en farmakologisk profil svarende til den i insekter.
Bjørnedyr kan overleve et bredt spektrum af osmotiske miljøer (semiterrestrisk,
limnisk og marine habitater), hvorfor evnen til at volumen- og osmoregulere
blev undersøgt. Disse studier demonstrerede at bjørnedyr kan regulere den
samlede kropsvolumen under såvel hypo- som hyperosmotiske forhold, samt
indikerede at hyperregulering kan være en gennemgående tendens hos
eutardigrader. Indeværende arbejder har altså bidraget til at fastslå bjørnedyr
som en vigtig eksperimentel gruppe, hvori centrale fysiologiske spørgsmål kan
besvares, heriblandt aspekter af ionregulering og osmoregulering.
29

Acknowledgements
During the course of my PhD studies I was fortunate to have the generous help
of many great people, which has made this work possible.
First and foremost, I thank my supervisor Nadja Møbjerg for being an
unwavering source of support, encouragement and guidance during past years,
but also for constantly challenging me to develop intellectually as well as a
researcher. I truly appreciate all you have done for me, and hope to be able to
continue our collaboration in the future.
I also greatly appreciate the support of Reinhardt Møbjerg Kristensen
(close to being my second supervisor) who initially introduced me to strange
world of tardigrades, and who took me under his wing, offering me guidance
and support (and equally important coffee). You have had enormous impact on
my education as a young researcher, and I have sincerely enjoyed working with
you.
I have had the great fortune of working with many wonderful
colleagues at University of Copenhagen, both at The August Krogh Center and
the Natural History Museum who all are warmly thanked. Thanks to you it has
been a pleasure coming to work every day. I am especially grateful to Erik
Hviid Larsen with whom I shared many interesting discussions, and who was
an endless source of inspiration. Aslak Jørgensen is thanked for great
collaborations and discussions over the years as well as for critical review of
my thesis. Dennis Krogh Persson is thanked for being a good colleague, but
more importantly, a good friend. Since we started this journey in research
together, I have come to depend on our collaborations and camaraderie, which
has been an invaluable help along the way (by the way, it is your turn to buy
cake). I thank Jette Lyby Michelsen for technical assistance over the years.
During my visits to Roskilde University I was fortunate to collaborate
with several great people. I especially thank Hans Ramløv for allowing me to
come work in his laboratory (on several occasions), which scientifically turned
out to be quite fruitful. I have particularly enjoyed our conversations and
discussions, and I thank you for both your encouragement and support. I also
thank Kristine Wulff Larsen for unparalleled project teamwork as well as our
many conversations on everything and anything. It was an absolute pleasure,
30

which I hope to repeat some time in the future. Peter Westh is thanked for
allowing me to work in his laboratory and for his enormous expertise and help.
Perhaps most importantly I thank the help and support of family and
friends. I especially thank my parents and parent in-laws for their invaluable
support during difficult times, and for helping me and my family in more ways
than I care to mention. You’re the best! Lastly I thank my girlfriend Iben Rønn
Veland for helping me, supporting me, tolerating me and loving me, but mostly
for taking care of our son when I was working – none of this was possible
without your help! I also thank my little son Hannibal for tolerating that I had
to work sometimes, and couldn’t be home when you wanted me to. I am
looking forward to making up for lost time. I love you both more than you
know. Thank you.
This work was funded by the 2008 Faculty of Science, University of
Copenhagen Freja-Programme.
31

References
Anderson, E. (1960). The ultramicroscopic structure of the reptilian kidney. J Morph 106:205-
240.
Aguinaldo, A.M.A., Turbeville, J.M., Linford, L.S., Rivera, M.C., Garey, J.R., Raff, R.A.
& Lake, J.A. (1997). Evidence for a clade of nematodes, arthropods and other moulting
animals. Nature 387 (6632):489-493.
Beyenbach, K. W. & Piermarini P. M. (2011). Transcellular and paracellular pathways of
transepithelial fluid secretion in Malpighian (renal) tubules of the yellow fever mosquito
Aedes aegypti. Acta Physiol 202:387-407.
Bertolani, R (2001). Evolution of the reproductive mechanisms in tardigrades – a review. Zool
Anz 240:247-252.
Campbell, L.I., Rota-stabelli, O., Edgecombe, G.D., Marchioro, T., Longhorn, S.J.,
Telford, M.J., Philippe, H., Rebecchi, L., Peterson, K.J. & Pisani, D. (2011).
MicroRNAs and phylogenomics resolve the relationships of Tardigrada and suggest that
velvet worms are the sister group of Arthropoda. PNAS 108:15920-15924.
Campiglia, S. (1976). The blood of Peripatus acacioi Marcus & Marcus (Onychophora) – III.
The ionic composition of the hemolymph. Comp Biochem Physiol 54(A):129-133.
Coast, G. (2007). The endocrine control of salt balance in insects. Gen Comp Endocrinol
152:332-338.
Crowe, J. H. & Madin, K. A. (1974). Anhydrobiosis in tardigrades and nematodes. Trans Am
Microsc Soc 93:513-524.
Dantzler, W. H. (2002). Renal organic anion transport: a comparative and cellular perspective.
Biochim Biophys Acta 1566:169-181.
Dawson, D. C. & Liu, Xuehong (2009). Osmoregulation: some principles of water and solute
transport. In: David H. Evans (ed) Osmotic and ionic regulation: cells and animals. CRS
Press, Florida, USA.
Degma, P., Bertolani, R. & Guidetti, R. 2009-2012. Actual checklist of Tardigrada species.
Ver. 20: 17-01-2012. http://www.tardigrada.modena.unimo.it/miscellanea/Actual checklist
of Tardigrada.pdf
Degma, P. & Guidetti, R. (2007). Notes to the current checklist of Tardigrada. Zootaxa,
1579:41-53.
Dewel, R. A. & Dewel, W. C. (1979). Studies on the tardigrades. J Morphol 161:79-110.
Diehl, W. J. (1986). Osmoregulation in echinoderms. Comp Biochem Physiol A 84:199-205.
Dow, J. A. T. & Davies, S. A. (2006). The Malpighian tubule: Rapid insights from postgenomic
biology. J Insect Physiol 52:365-378.
Dowd, W. W., Harris, B. N. Chech Jr, J. J. & Kültz, D (2010). Proteomic and physiological
responses of leopard sharks (Triakis semifasciata) to salinity change. J Exp Biol 213:210-
224.
32

Dunn, C.W., Hejnol, A., Matus, D.Q., Pang, K., Browne, W.E., Smith, S.A., Seaver, E.,
Rouse, G.W. Obst, M., Edgecombe, G.D., Sørensen, M.V., Haddock, S.H.D.,
Schmidt-Rhaesa, A., Okusu, A., Kristensen, R.M. & Wheeler, W.C. (2008). Broad
phylogenomic sampling improves resolution of the animal tree of life. Nature 452:745-
750.
Edgecombe, G. D. (2010). Arthropod phylogeny: An overview from the perspective of
morphology, molecular data and the fossil record. Arthropod Struct Dev 39:74-87.
Essig, A. (1968). The “pump-leak“ model and exchange diffusion. Biophys J 8(1):53-63.
Gabriel, W. N. & Goldstein, B. (2007). Segmental expression of Pax3/7 and Engrailed
homologs in tardigrade development. Dev Genes Evol 217:421-433.
George, R. L., Wu, X., Huang, W., Fei, Y. J., Leibach, F. H. & Ganapathy, V. (1999).
Molecular cloning and functional characterization of a polyspecific organic anion
transporter from Caenorhabditis elegans. J Pharmacol Exp Ther 291:596-603.
Gilles, R. & Delpire, E. (1997). Variations in salinity, osmolarity and water availability. In:
Dantzler, W. H. (ed.). Handbook of comparative physiology. New York: Oxford
University Press, USA.
Glasheen, J. S. & Hand, S. C. (1988). Anhydrobiosis in embryos of the brine shrimp Artemia:
characterization of metabolic arrest during reductions in cell associated water. J Exp Biol
135:363-380.
Goldstein, B. & Blaxter, M. (2002). Tardigrades. Curr Biol 12:R475.
Guidetti, R. & Bertolani, R. (2005). Tardigrade taxonomy: an updated check list of the taxa
and a list of characters for their identification. Zootaxa 845:1-46.
Guidetti, R., Bertolani, R. & Nelson, D. R. (1999). Ecological and faunistic studies on
tardigrades in leaf litter of beech forest. Zool Anz 238:215-223.
Hallas T, Yeates G (1972) Tardigrades of the soil and litter of a Danish beech forest.
Pedobiologia 12:287-304.
Hincha, D. K. & Hagemann, M. (2004). Stabilization of model membranes during drying by
compatible solutes involved in the stress tolerance of plants and microorganisms. Biochem
J 383:277-283.
Hobsen, A. D., Stephenson, W. & Eden A. (1952). Studies on the physiology of Ascaris
lumbricoides: II. The inorganic composition of the body fluid in relation to that of the
environment. J Exp Biol 29:22-29.
Jönson, K. I. (2003). Population density and species composition of moss-living tardigrades in
a boreo-nemoral forest. Ecography 26:356-364.
Jørgensen, A., Faurby, S., Hansen, J. G., Møbjerg, N. & Kristensen, R. M. (2010).
Molecular phylogeny of Arthrotardigrada (Tardigrada). Mol Phylogenet Evol 54:1006-
1015.
Jørgensen, A., Møbjerg, N. & Kristensen, R. M. (2007). A molecular study of the tardigrade
Echiniscus testudo (Echiniscidae) reveals low DNA sequence diversity over a large
geographical area. Proceedings of the Tenth Internationl Symposium on Tardigrada. J
limnol 66:77-83.
Liggins, G. W. & Grigg, G. C. (1985). Osmoregulation of the cane toad Bufo marinus, in salt
water. Comp Biochem Physiol A 82:613-619.
Macallum, A. B. (1910). The inorganic composition of the blood in vertebrates and
invertebrates, and its origin. Proc R. Soc Lond B 82(559):602-624.
33

Maddrell, S. H. P. & Phillips, J. E. (1975). Secretion of hypo-osmotic fluid by the lower
Malpighian tubules of Rhodnius prolixus. J Exp Biol 62:671-683.
Martinez, E. A. (1975). Marine meiofauna of a New York City beach, with particular
reference to Tardigrada. Est Coast Mar Sci 3:337-348.
Morgan, C. I. (1977). Population dynamics of two species of Tardigrada, Macrobiotus
hufelandii (Schultze) and Echiniscus (Echiniscus) testudo (Doyère), in roof moss from
Swansea. J Anim Ecol 46:263-279.
Møbjerg, N & Dahl, C. (1996). Studies on the morphology and ultrastructure of the
Malpighian tubules of Halobiotus crispae Kristensen, 1982 (Eutardigrada). Zool J Linn
Soc 116:85-99.
Møbjerg, N., Jespersen, Å & Wilkinson, M. (2004). Morphology of the kidney in the West
African caecilian, Geotrypetes seraphini (Amphibia, Gymnophiona, Caeciliidae). J Morph
262(2):583-607.
Nelson, D. R. (2002). Current status of Tardigrada: evolution and ecology. Integr Comp Biol
42(3):652-659.
Normant, M., Kubicka, M., Lapucki, T., Czarnowski, W. & Michalowska M. (2005).
Osmotic and ionic haemolymph concentration in the Baltic Sea amphipod Gammarus
oceanicus in relation to water salinity. Comp Biochem Physiol A 141:94-99.
O’Donnell, M. J. & Maddrell, S. H. P. (1995). Fluid reabsorption and ion transport by the
lower Malpighian tubules of adult female Drosophila. J Exp Biol 198:1647-1653.
Peaker, M (1971). Avian salt glands. Philos Trans R Soc Lond B 262:289-300.
Pelzer, B., Dastych, H. & Greven, H. (2007). The osmoregulatory/excretory organs of the
glacier-dwelling eutardigrade Hypsibius klebelsbergi Mihelcic, 1959 (Tardigrada). Mitt.
hamb. Zool Mus Inst 104:61-72.
Ramazzotti, G. & Maucci W. (1983). Il Phylum Tardigrada. Terza edizione riveduta e
corretta. Mem Insti Ital Idro Dott Marco De Marchi 41:1-1012.
Reilly, D., Cramp, R. L., Wilson, J. M., Campbell, H. A. & Franklin, C. E. (2011).
Branchial osmoregulation in the euryhaline bull shark, Charcharhinus leucas: a molecular
analysis of ion transporters. J Exp Biol 214:2883-2895.
Renaud-Mornant, J. (1982). Species diversity in marine Tardigrada. In: Nelson (Ed.)
Proceedings of the 3rd international symposium on Tardigrada. East Tennessee State
University Press, pp. 149-178.
Reynolds, S. E. & Bellward, K. (1989). Water balance in Manduca sexta caterpillars: water
recycling from the rectum. J Exp Bio 141:33-45.
Ricci, C., Melone, G. Santo, N. & Caprioli, M. (2003). Morphological response of a Bdelloid
Rotifer to dessication. J Morphol 257:246-253.
Riegel, J. A. (1970). A new model of transepithelial fluid movement with detailed application
to fluid movement in the crayfish antennal gland. Comp Biochem Physiol 36:403-410.
Robertson, J. D. (1949). Ionic regulation in some marine invertebrates. J Exp Biol 26:182-
200.
Rota-Stabelli O, Kayal E, Gleeson D, Daub J, Boore J, Telford M, Pisani D, Blaxter M,
Lavrov D. (2010). Ecdysozoan mitogenomics: evidence for a common origin of the
legged invertebrates, the Panarthropoda. Genome Biol Evol 2:425-440.
Sardella, B. A., Baker, D. W. & Brauner, C. J. (2009). The effects of variable water salinity
and ionic composition on the plasma status of the Pacific Hagfish (Eptatretus stoutii). J
Comp Physiol B 179:721-728.
34

Sarver, R. G., Flynn, M. A. & Holliday, C. W. (1994). Renal Na, K-ATPase and
osmoregulation in the crayfish Procambarus clarkii. Comp Biochem Physiol (A) 107:349-
356.
Schluter, D. (2009). Evidence for ecological speciation and its alternative. Science 323:737-
741.
Schmidt-Nielsen, K. (1963). Osmotic regulation in higher vertebrates. Harvey lect 58:53-93.
Schmidt-Rhaesa A, Kulessa J. (2007). Muscular architecture of Milnesium tardigradum and
Hypsibius sp. (Eutardigrada, Tardigrada) with some data on Ramazzottius oberhaeuseri.
Zoomorphol 126:265-281.
Sutcliffe, D. W. (1962). The composition of haemolymph in aquatic insects. J Exp Biol
39:325-343.
Suzuki, A. C. (2003). Life history of Milnesium tardigradum Doyère (Tardigrada) under a
rearing environment. Zool Sci 20:49-57.
Torrie, L. S., Radford, J. C., Southall, T. D., Kean, L., Dinsmore, A. J., Davies, S. A. &
Dow, J. A. T. (2004). Resolution of the insect ouabain paradox. PNAS 101: 13689-13693.
Uhía, E. & Briones, M. J. I. (2002) Population dynamics and vertical distribution of
enchytraeids and tardigrades in response to deforestation. Acta Oecolog 23:349-359.
Van Weel, P. B. (1957). Observations on the osmoregulation in Aplysia juliana Pease
(Aplysiidae, Mollusca). Z vergl Physiol 39:492-506.
Walz, B. (1974). The Fine Structure of Somatic Muscles of Tardigrada. Cell Tiss Res 149:81-
89.
Weglarska, B. (1987). Studies on the excretory system of Isohypsibius granulifer Thulin
(Eutardigrada). In: Biology of tardigrades. Selected symposia and monographs (1) (ed.
Bertolani, R.), pp. 15-24. Modena: U.Z.I. Mucchi.
Whittamore, J. M. (2012). Osmoregulation and epithelial water transport: lessons from the
intestine of marine teleost fish. J Comp Physiol B 182:1-39.
Wilder, M. N., Ikuta, K., Atmomarsono M., Hatta, T. & Komuro, K. (1998). Changes in
osmotic and ionic concentrations in the hemolymph of Macrobrachium rosenbergii
exposed to varying salinities and correlation to ionic and crystalline composition of the
cuticle. Comp Biochem Physiol A 119:941-950.
Williams, J. C. & Beyenbach, K. W. (1983). Differential effects of secretagogues on Na and
K secretion in the Malpighian tubules of Aedes aegypti. J Comp Physiol B 149:511-517.
Yancey, P. H. (2005). Organic osmolytes as compatible, metabolic and counteracting
cytoprotectants in high osmolarity and other stresses. J Exp Biol 208:2819-2830.
Zantke, J., Wolff, C. & Scholtz, G. (2008). Three-dimensional reconstruction of the central
nervous system of Macrobiotus hufelandi (Eutardigrada, Parachela): implications for the
phylogenetic position of Tardigrada. Zoomorphol 127:21–36.
Zhao, H. (2005). Effect of ions and other compatible solutes on enzyme activity, and its
implication for biocatalysis using ionic liquids. J Mol Catal B: Enzymat 37:16-25.
35

36
UNIVERSITAS
HAFNIENSIS
2012

Paper I

Inorganic ion composition in Tardigrada: cryptobionts contain large
fraction of unidentified organic solutes
Kenneth Agerlin Halberg 1,* , Kristine Wulff Larsen 2 , Aslak Jørgensen 3 , Hans Ramløv 2 and
Nadja Møbjerg 1
1 Department of Biology, the August Krogh Centre, University of Copenhagen, Universitetsparken 13, DK-
2100 Copenhagen Ø, Denmark, 2 Department of Nature, Systems and Models, University of Roskilde,
Universitetsvej 1, DK-4000 Roskilde, Denmark and 3 Laboratory of Molecular Systematics, Natural History
Museum of Denmark, University of Copenhagen, Sølvgade 83, DK-1307 Copenhagen K, Denmark
*Author for correspondence (kahalberg@bio.ku.dk)
Submitted June 2012
SUMMARY
Tardigrades are a group of micrometazoans known to tolerate extreme environmental stress. Significant
efforts have been devoted to the field, however; mechanisms explaining the extreme adaptations found
among tardigrades is still lacking. Here we present data on the inorganic ion composition and total
osmotic concentration of five different species of tardigrades (E. testudo, M. tardigradum, R. coronifer, M.
cf. hufelandi and H. crispae) using high-performance anion-exchange chromatography and nanoliter
osmometry. Quantification of the ionic content indicates that Na + and Cl - are the principle inorganic ions
in tardigrade fluids, albeit substantial concentrations of K + , NH + 4 , Ca 2+ , Mg 2+ , F - 2-
, SO 4 and PO 3- 4 also
were detected. In limno-terrestrial tardigrades, the respective ions are concentrated by a large factor
compared to that of the external medium (Na + , ×70-800; K + , ×20-90; Ca 2+ and Mg 2+ , ×30-200; Cl - , ×20-50;
SO 2- 4 , ×30-150), whereas in the marine species H. crispae Na + , Cl - 2-
and SO 4 are almost in ionic
equilibrium with (brackish) salt water, while K + , Ca 2+ and Mg 2+ are slightly concentrated (×2-10).
However, there is an anion deficit of ~120 mEq/l in M. tardigradum and H. crispae, indicating that there
are ionic components that remain unidentified in these species. Body fluid osmolality ranged from 361±49
in R. coronifer to 961±43 mOsm/kg in H. crispae. Concentrations of most inorganic ions are largely
identical between active and dehydrated groups of R. coronifer, suggesting that this tardigrade does not
exclude large amounts of ions during dehydration. The large osmotic and ionic gradients maintained by
both limno-terrestrial and marine species are indicative of a powerful ion-retentive mechanism in
Tardigrada. Moreover, our data indicates that cryptobiotic tardigrades contain a large fraction of
unidentified organic osmolytes, the identification of which is expected to provide increased insight into
the phenomenon of cryptobiosis.
Key words: tardigrades, inorganic ions, ion chromatography, nanoliter osmometry, organic osmolytes,
cryptobiosis
INTRODUCTION
Tardigrades are a group of minute multi-cellular
animals that are known to tolerate extreme
environmental stress (Guidetti et al., 2010;
Møbjerg et al., 2011). This capacity derives
mainly from their ability to enter a state latent of
life, i.e. cryptobiosis, in which their resistances to
adverse environmental conditions are greatly
increased (Møbjerg et al., 2011). In recent years,
intensive research efforts have been devoted to the
field, as the translational output associated with a
detailed understanding of their complex stress
biology is expected to include new methods for
preserving and stabilizing biological materials
(Wełnicz et al., 2010). Several advances have been
made in our understanding of tardigrade stress
responses, especially regarding i) the role of
selective carbohydrates (trehalose), ii) the

K. A. Halberg and others
differential expression of stress proteins (heat
shock proteins and late embryogenesis abundant
proteins), and iii) the identification of antioxidant
defenses and DNA repair mechanisms
(see reviews by Guidetti et al., 2010; Wełnicz et
al., 2010; Møbjerg et al., 2011). However, a
general mechanism explaining the extreme
adaptations found among tardigrades is still
lacking. Consequently, new approaches that may
provide increased insight into the superior stress
adaptations of tardigrades are greatly needed.
Knowledge of the composition as well as
concentrations of dissolved particles in internal
fluids is fundamental to the understanding of
basic physiological processes, such as fluid and
electrolyte homeostasis, signal transduction and
solute transport. Accordingly, such data have
been provided for most major groups of animals
(cnidarians, echinoderms, annelids, molluscs,
crustaceans, insects, chelicerates, tunicates, fish,
amphibians and mammals) more than half a
century ago (Macallum, 1910; Robertson, 1949,
1954; Sutcliffe, 1962; Hronoski & Armstrong,
1977). However, practically nothing is known
about the chemical composition of tardigrades,
which has been a major obstacle to the
understanding of the fluid and solute dynamics in
these animals (Halberg et al., 2009b; Møbjerg et
al., 2011; Halberg & Møbjerg, 2012). Questions
relating to this area of research are especially
important to address, if we wish to unravel the
biological mechanisms mediating the unique
tolerance to extreme desiccation (anhydrobiosis)
– the most widespread form of cryptobiosis in
Tardigrada.
In the present study, we use a combination of
high-performance anion-exchange chromatography
(HPAEC) and nanoliter osmometry, to
identify and quantify inorganic cations and anions
present in tardigrade homogenates, and to
measure the total osmotic concentrations of five
different species of tardigrades, covering both a
broad phylogenetic and habitat spectrum. Our
study indicates that tardigrades possess powerful
ion-retentive and osmoregulatory capacities, and
that (only) cryptobiotic species contain a large
fraction of organic solutes.
MATERIALS AND METHODS
Tardigrade sampling
Specimens of Richtersius coronifer Richters,
1908, Macrobiotus cf. hufelandi C.A.S. Schultze,
1834 and Milnesium tardigradum Doyère, 1840
were extracted from moss collected at Öland,
Sweden, while Echiniscus testudo Doyère, 1840
was found in moss collected at Nivå, Denmark.
These species were extracted by washing the
respective moss samples with tap water through
six different sieves of progressively smaller mesh
size, so as to concentrate the tardigrades and to
remove large debris. Specimens of Halobiotus
crispae Kristensen, 1982 were isolated from
marine algae and sediment collected from
Vellerup Vig, Denmark according to the method
of Halberg and Møbjerg (2012). The total number
of animals used in all experiments was 2220 R.
coronifer, 326 M. tardigradum, 630 M. cf.
hufelandi, 426 E. testudo and 268 H. crispae.
Inorganic cation and anion analysis
The dominant cations and anions present in the
different tardigrade species were determined by
HPAEC using a Metrohm chromatography system
(830 IC interface, 818 IC pump, 819 IC
conductivity detector, columns C4 - 150/4.0
(cations) and A supp 5 150/4.0 (anions); Metrohm,
Herisau, Switzerland). The eluents (mobile
phases) were made according to manufacturer’
instructions: For cations, the eluent consisted of
0.7 mM C 7 H 5 NO 4 + 1.7 mM (65%) HNO 3 . For
anions, it consisted of 3.2 mM Na 2 CO 3 + 1.0 mM
NaHCO 3 . The eluents were filtered (mesh size: 45
µm) prior to use. The analysis settings employed
were a flow-rate of 0.9 ml/min (cations) and 0.7
ml/min (anions) with a pressure of ~6.4 MPa.
Cation analyses were performed non-suppressed,
whereas anion detection was conducted using
chemical suppression. Fluka multi-element cation
and anion standards (Sigma-Aldrich, St. Louis,
MO, USA) were used to construct calibration
curves for the respective ions bracketing the
concentration range of interest. Based on these
calibration curves, the ion chromatography (IC)
software (IC Net 2.3, Metrohm, Herisau,
Switzerland) calculated the ion concentrations of
all subsequent samples (mg/l), which were
recalculated to a different unit of concentration
(mM) and adjusted according to the appropriate
dilution factor (see below). Representative
chromatograms of both the cationic and anionic
fractions are shown for all investigated species
(Figs. 1, 2). The empirically determined elution
order and retention times of the investigated ions
were Na + (t R = 5.37 min), NH 4 + (t R = 6.03 min),

Inorganic ion composition in Tardigrada
Fig. 1. Representational chromatograms revealing the principal inorganic cations present in each investigated species:
1, sodium; 2, ammonium; 3, potassium; 4, calcium; 5, magnesium; nOAp, negative organic acid peak. Stitched square
indicates an unidentified compound (t R = 10.36 min) that increases app. two-fold in absolute concentration in
dehydrated animals of Richtersius coronifer (data not shown). Column, Metrohm C4-150/4.0; mobile phase, 0.7 mM
dipicolinic acid + 1.7 mM (65%) nitric acid; flow-rate, 0.9 ml/min; Conductivity detector without suppression.
Injection volume, 60 µl.
K + (t R = 7.73 min), Ca 2+ (t R = 18.12 min), Mg 2+
(t R = 23.17 min), F - (t R = 4.05 min), Cl - (t R = 6.01
min), PO 4 3- (t R = 14.74 min) and SO 4 2- (t R = 16.58
min).
Sample preparation
Following extraction, specimens were washed
repeatedly with ddH 2 O (Halobiotus crispae was
washed in filtered salt water; SW, 20 ‰), and
subsequently transferred, using an Irwin loop, to
sample tubes containing cation eluent (75-100
µl); samples dissolved in cation eluent allows for
a more precise quantification of cations due to
increased signal to noise ratio (pers. comm.;
Metrohm Nordic, Denmark). Prior to transfer,
surface water was removed by blotting the
animals with tissue paper in an attempt to avoid
unwanted dilution of the samples. Pilot
experiments revealed that this process was critical
to acquire reproducible data, and was accordingly
performed as fast and uniform as possible. A total
of 40-225 animals were transferred to each test
tube and the sample was subsequently
homogenized using a sterile plastic pestle; great
care was taken to ensure complete homogenization
(visually confirmed at 50× magnification), and the
pestle was subsequently rinsed with a small
volume of cation eluent to ensure total transfer of
ions to the test tube. The number of animals per
sample (N) varied according to species size and
availability (Tables 1, 2). The entire sample was
then centrifuged (10 min at 5600 rpm) to remove
solid particles (e.g. cuticle fragments), and the
supernatant was filtered (mesh size: 0.20 µm)
using a single use syringe filter (Sartorius AG,
Göttingen, Germany). The samples were
subsequently frozen at -20 ºC, if not quantified
immediately. A total of five to seven samples were
prepared for each species (Table 2).

K. A. Halberg and others
Fig. 2. Representational chromatograms revealing the principal inorganic anions present in each investigated species:
6, fluoride; 7, chloride; 8, phosphate; 9, sulfate; W, negative water peak. Stitched square indicates an unidentified
compound (t R = 7.24 min) that increases app. two-fold in absolute concentration in dehydrated animals of Richtersius
coronifer (data not shown). Column, Metrohm A supp 5 150/4.0; mobile phase, 3.2 mM sodium carbonate + 1.0 mM
sodium hydrogen carbonate; flow-rate, 0.7 ml/min; Conductivity detector with chemical suppression. Injection
volume, 60 µl.
The ionic concentration and composition of
the external media from the different habitats i.e.
moss water (MW) and SW, were additionally
determined. Moss samples were rehydrated in
ddH 2 O for several hours, and MW samples were
subsequently collected from between the leafcovered
stems. SW samples were prepared by
diluting SW (1:200) collected at the locality.
Samples were quantified in triplets using both
vapor pressure osmometry (Vapro 5520, Wescor
inc., UT, USA) and HPAEC (Table 3).
In order to document whether changes in
inorganic ion content occur during dehydration
from an active to a cryptobiotic state, samples of
dehydrated Richtersius coronifer were
additionally prepared. Groups of 75 animals were
transferred to each sample tube, and excess water
was removed by blotting the animals with tissue
paper. The animals were subsequently allowed to
dehydrate over the ensuing 24 h at ambient
temperature and humidity. Following complete
dehydration, tissue paper saturated with ddH 2 O
was used to rinse the surface of the animals, and a
dry tissue paper was used to remove excess
moisture. This was done in order to remove
potential solutes extruded on the surface of the
animals – the surface of some animals was
inaccessible due to animal clumping, and therefore
could not be rinsed. A volume of 90 µl of cation
eluent was added, and the animals were
immediately homogenized. The samples were then
prepared as described above with six samples
prepared in total (Table 2). Data (mg/l) from this
experiment was directly compared to that of
hydrated animals (Table 4; Fig. 3B), as both sets
of samples contained near identical number of
animals per unit volume (i.e. 0.81 and 0.83
animals/µl eluent respectively), which
circumvented the need for recalculations (see
below). In order to test whether R. coronifer
actually produced viable tuns during the
abovementioned conditions, post-cryptobiotic

Inorganic ion composition in Tardigrada
survival was assessed. Using four groups of 50
specimens, a survival rate of 94 ± 4% (mean ±
s.d.) was observed, which is comparable to the
maximally reported survival rate of R. coronifer
dehydrated on Whatman filters (Persson et al.,
2010).
Calculation of ion concentrations
The IC software expressed the integration of
peaks as a concentration (mg/l), which was
recalculated using the respective molecular
weights of each compound to a different
concentration (mM) prior to adjusting for the
dilution factors.
The volume of each investigated species was
calculated according to an adjusted method of
Halberg et al. (2009b). In brief, micrographs were
taken of N = 20 animals of each species, using a
digital camera (C-5050, Olympus, Japan)
mounted on an Olympus BX 51microscope
(Olympus, Japan), and median length (h) and
width (2r) of the trunk and legs were measured.
Approximating the geometric shape of the trunk
and legs as a cylinder, and adjusting the volume
of liquid according to the gravimetrically
measured water content (based on Westh and
Kristensen, 1992; Halberg et al., 2009b), the fluid
volume of an individual tardigrade of each
species was calculated using equation (1).
(Eq 1) V individual = π(r 2 trunkh trunk + 8r 2 legh leg ) × W
Where V individual is the volume of an individual
tardigrade, r is the radius and h the length of the
trunk and hind legs respectively, while W (0.72 i.e.
mean fractional water content of R. coronifer and
H. crispae) is the gravimetrically measured
fractional water content. Using these data (Table
1), the total tardigrade test volume was calculated
by multiplying the volume of an individual with
the number of animals included in the sample
according to equation (2).
(Eq 2) V total = V individual × N
V total is the total tardigrade test volume, and N the
number of animals included in the sample. Lastly,
the concentrations of the dominant cations and
anions in the investigated species of tardigrades
were calculated by multiplying the measured ion
concentrations with the dilution factor, which was
calculated according to equation (3).
(Eq 3) D = F / V total
Where D is the dilution factor and F is the final
volume (i.e. volume of cation eluent the
tardigrades were transferred to + V total ). Sample
information for the respective species is listed in
Table 2.
Nanoliter osmometry
The total osmotic concentration of tardigrades
from each investigated species was estimated
using nanoliter osmometry. This was done in order
to determine the fraction that the identified
Table 1. Volume estimations. Mean values of length (h) and width (2r) of the trunk and legs (N = 20 animals), as well
as the calculated volume (Eq 1), of each investigated species. W is the average of the gravimetrically determined
water content (72%) of Richtersius coronifer (Westh and Kristensen, 1992) and Halobiotus crispae (Halberg et al.,
2009). Data are expressed as mean ± s.d.

K. A. Halberg and others
inorganic ions constitute of the total osmotic
concentration in each species, and to provide an
independent verification of our HPAEC data, i.e.
total osmotic concentration should be higher than
the accumulated concentration of the respective
inorganic ions (Table 3). Using the same
procedure for removing excess water as described
above, individual specimens were transferred into
sample oil wells (loading oil type B; cST=1250 ±
10%; Cargille laboratories, Cedar grove, NJ
07009, USA) of a calibrated nanoliter osmometer
(Clifton Technical Physics, Hartford, NY, USA),
and the osmolality (mOsm/kg) was determined by
freezing point depression (FPD = 1.858
°C/Osmol). Six to ten animals of each species
were used in this experiment (Table 3).
Statistics
Significant changes in the individual inorganic
ion concentrations between active and
cryptobiotic animals of Richtersius coronifer
were tested using an unpaired, two-sample t-test
with significance levels of P>0.05 (not
significant, NS), P

Inorganic ion composition in Tardigrada
Figure 3. Graphical representation of the respective ionic contributions to total osmotic concentration. A.
Concentrations (mM) of the respective cations and anions measured in each investigated species, as well as the
corresponding total osmotic concentration (mOsm/kg), as measured by nanoliter osmometry (see also Table 3). The
blank area represents the osmotic deficit (OD), i.e. other solutes. The phylogenetic position and habitat preference of
each species is listed. The light micrographs of the animals are shown to scale (see Table 1 for average species size).
B. Concentrations (mg/l) of the respective cations and anions measured in hydrated, active specimens compared to
dehydrated cryptobiotic animals of Richtersius coronifer (see also Table 4). Data are expressed as mean ± s.d.

K. A. Halberg and others
Halobiotus crispae containing the lowest and
highest concentrations respectively (Fig. 3A;
Table 3). There are notable differences in the Na + /
Cl - ratio between the animals, i.e., the ratio is less
than unity in the limno-terrestrial herbivores E.
testudo (0.61), R. coronifer (0.60) and M. cf.
hufelandi (0.33), higher than unity in the limnoterrestrial
predator M. tardigradum (1.19) and
close to unity in the marine herbivore H. crispae
(0.92).
Compared with Na + and Cl - , generally, the
[K + ] is relatively low in all species, ranging from
19-73 mM (Tab. 3). Thus, the Na + / K + ratio is
higher than unity in E. testudo (2.39), M.
tardigradum (2.05), R. coronifer (1.32) and H.
crispae (6.12), however; lower than unity in M.
cf. hufelandi (0.77). Interestingly, the relative
contribution of K + to total osmotic concentration
is low constituting

Inorganic ion composition in Tardigrada
Table 3. Ionic composition and total osmotic concentration of the investigated species of tardigrades and the corresponding external media. Concentrations (mM) of
cations and anions detected in each investigated species, as well as moss water (MS) and 20 ‰ salt water (SW) samples, in addition to the corresponding total osmotic
concentration (mOsm/kg), as measured by nanoliter osmometry or vapor pressure osmometry, respectively. In addition, the osmotic deficits (calculated as the
difference in ionic concentration, and total osmotic concentration), as well as the observed charge deficits (calculated as the difference between positive and negative
charges) are listed; the polarity of the charge deficits is indicated in parenthesis. Numbers noted in brackets indicates the number of samples tested. Data are expressed
as mean ± s.d.

K. A. Halberg and others
In the present study we provide data on the ionic
composition of five different species of
tardigrades covering a large phylogenetic
spectrum. Our study is represented by members
of Heterotardigrada (Echiniscoidea) and
Eutardigrada (Apochela and Parachela), four
evolutionary distant families (Echiniscidae,
Milnesiidae, Macrobiotidae and Hypsibiidae), as
well as both limno-terrestrial and marine habitats.
Accordingly, we will discuss the ionic
compositions of the respective species in relation
to systematic position and habitat preference, as
well as make comments on our data in relation to
hemolymph composition in representatives of
phylogenetically related groups (i.e. Arthropoda
and Onychophora).
Echiniscus testudo (Heterotardigrada:
Echiniscidae) belongs to another evolutionary
lineage than the other tardigrades in the present
study. Compared to limno-terrestrial members of
Eutardigrada, the ionic composition of this
heterotardigrade is characterized by a large
contribution of Na + and Cl - (~45%), and a very
low contribution of Mg 2+ 2-
(0.4%), SO 4 (0.6%)
and PO 3- 4 (1.4%), respectively (Tab. 5). The large
contribution of Na + and Cl - to total osmotic
concentration, which is comparable to that seen in
the marine species H. crispae (Table 5), could
reflect the supposed marine origin of tardigrades
(Jørgensen et al., 2010). This hypothesis can be
tested by data on members of the ‘ancient’ and
exclusively marine Arthrotardigrada.
The family Milnesiidae, represented by the
predator Milnesium tardigradum, is currently
considered the sister-group of all other
eutardigrades (Guidetti et al., 2009). M.
tardigradum contains the highest total osmotic- as
well as ionic concentration among the limnoterrestrial
species, with conspicuously high levels
of both K + and Ca 2+ (Table 3). The high [K + ] in
M. tardigradum compared to the phytophagous
species is somewhat surprising, as e.g.
carnivorous insects typically contain low levels of
K + (Sutcliffe, 1962). Conversely, phytophagous
tardigrades are known to feed on bryophytes high
in K + and low in Na + (Smith, 1978), and were,
analogous to phytophagous insects (Sutcliffe,
1962), expected to reflect this relative ion
composition in their extracellular body fluids.
Interestingly, the relative ion contributions to
total osmotic concentration suggests that M.
tardigradum resembles the heterotardigrades more
than the other eutardigrades (Table 5)
Sutcliffe (1962, 1963) argued that definitive
types of hemolymph are related to phylogenetic
position within Insecta. Apart from small
differences in absolute concentrations of Na + and
PO 3- 4 , the ionic compositions and relative
contributions of the different components in
Richtersius coronifer and Macrobiotus cf.
hufelandi (Eutardigrada: Macrobiotidae) are
similar (Tables. 3 and 5). As a testable hypothesis,
these similarities would suggest that the relative
ion composition among the species relates to
phylogeny and systematic position in Tardigrada.
In contrast to the other groups of limno-terrestrial
tardigrades, the inorganic content of R. coronifer
and M. cf. hufelandi is characterized by a
relatively small contribution of Na + (~2-7 times
lower) and Cl - (~2-3 times lower), and conversely,
2-
a relatively large contribution of SO 4 (~4-10
times higher). The physiological significance of
these variations is unknown.
Halobiotus crispae (Eutardigrada:
Hypsibiidae) is a truly marine species, and is the
species with the highest total concentration of both
ions and total solutes measured. Na + and Cl -
account for more than 50% of its total osmotic
concentration. The divalent cations, Ca 2+ and
Mg 2+ , are also detected in high concentrations,
both absolute and relative. In contrast to the
limno-terrestrial species, the total osmotic
concentration of H. crispae is almost exclusively
accounted for by the measured ionic
concentrations, which becomes evident when
considering the charge deficit indicated in Table 3.
The contribution of the total diffusible ions to
the total osmotic concentration of tardigrades is
roughly similar to that of the hemolymph of
arthropods, nematodes and onychophorans (Table
5). In fact, as Na + predominantly is an
extracellular ion, whereas K + and Ca 2+ mainly are
intracellular ions, the ionic composition of
tardigrade hemolymph is expected to resemble
that of closely related animal groups, e.g. other
members of Panarthropoda (see Table 5).
It is relevant to compare concentrations of ions
in tardigrade body fluids with those of the
respective external media (Table 3). The strongest
ability to concentrate ions is seen in limnoterrestrial
species, which hyper-regulate by as
much as ~350-750 mOsm/kg (Fig. 3A). The
marine H. crispae maintains hemolymph osmotic

Inorganic ion composition in Tardigrada
Table 4. Changes in ionic
composition during dehydration.
Concentrations (mg/l) of
the respective cations and
anions measured in hydrated,
active specimens compared to
dehydrated cryptobiotic animals
of Richtersius coronifer. Both
sets of samples contained near
identical number of animals per
unit volume (i.e. 0.8 animals/µl
eluent), and were accordingly
directly comparable. Numbers
noted in brackets indicates the
number of samples tested. Data
are expressed as mean ± s.d.
Significant difference in the
concentration of the respective
inorganic ion concentrations
were tested using an unpaired,
two-sample t-test with
significance levels of P>0.05
(not significant, NS), P

K. A. Halberg and others
intracellular osmotic potential so that osmotic
equilibrium between intra- and extracellular
fluids is maintained. Moreover, organic
osmolytes are known to stabilize macromolecular
structures by direct interaction with proteins and
membrane lipids (Crowe et al., 1987; Hincha and
Hagemann, 2004; Yancey, 2005). As evidenced
by the large osmotic deficits, which appears
restricted to cryptobiotic tardigrades (Fig. 3A;
Table 3), it is tempting to suggest that a large
quantity of organic osmolytes are synthesized in
these species, thus enabling the animals to
respond quickly to decreases in external water
potential. Such a strategy seems favorable in light
of the continuous dehydration-rehydration cycles
that may occur in limno-terrestrial habitats,
additionally supported by the short time span
(

Table 5. Total osmotic concentration (mOsm/kg) of the external medium and internal body fluids, as well as the osmotic contribution (%) of the respective ions to the
internal concentration in each of the investigated species of tardigrades. Corresponding data on hemolymph concentration and composition of selected species of
nematodes, crustaceans, insects and onychophorans is included for comparative purposes. TR, terrestrial; ─, not measured.
Inorganic ion composition in Tardigrada

K. A. Halberg and others
We would like to thank Anne Lise Maarup for technical
assistance and Reinhardt Møbjerg Kristensen (Natural History
Museum of Denmark) for loan of the Olympus BX 51
microscope. Station Linné (Porten til Alvaret), Ölands Skogsby,
Sweden is warmly thanked for accommodation during sampling
of tardigrades. Funding came from the Carlsberg Foundation
and the Freja-Programme (Faculty of Science, University of
Copenhagen).
FPD
HPAEC
IC
MW
SW
LIST OF ABBREVIATIONS
freezing point depression
high-performance anion-exchange chromatography
ion chromatography
moss water
salt water
REFERENCES
Campiglia, S. (1976). The blood of Peripatus acacioi
Marcus & Marcus (Onychophora) – III. The ionic
composition of the hemolymph. Comp. Biochem.
Physiol., 54(A):129-133.
Chen, T., Acker, J. P., Eroglu, A. flere authors
(2001). Beneficial effect of intracellular trehalose on
the membrane integrity of dried mammalian cells.
Cryobiol. 43:168-181.
Crowe, J. H., Crowe, L. M., Carpenter, J. F. and
Wistrom, C. A. (1987). Stabilization of dry
phospholipid bilayers and proteins by sugars.
Biochem. J., 242:1-10.
Bird, A. F., McClure, S. G. (1997). Composition of the
stylets of the tardigrade, Macrobiotus cf.
pseudohufelandi. Trans R Soc. S Aust., 121:43-50.
Eagers, R. Y. (1969). Toxic properties of inorganic
fluoride compounds. Elsevier, Amsterdam London
New York, 382 pp.
Granat, L. (1972). On the relation between pH and the
chemical composition in atmospheric precipitation.
Tellus XXIV, 6.
Guidetti, R., Altiero, T. & Rebbecchi, L. (2010). On
dormancy strategies in tardigrades. J. Insec.
Physiol., 57(5):567-576.
Guidette, R., Schill, R. O., Bertolani, R., Dandekar
T. & Wolf, M. (2009). New molecular data for
tardigrade phylogeny, with the erection of
Paramacrobiotus gen. nov. J. Zool. Syst. Ecol. Res.,
47(4):315-321.
Halberg, K. A., Persson D., Møbjerg N., Wanninger
A. & Kristensen R. M. (2009a). Myoanatomy of
the Marine Tardigrade Halobiotus crispae
(Eutardigrada: Hypsibiidae). J. Morphol., 270:996-
1013.
Halberg, K. A., Persson, D., Ramløv, H., Westh, P.,
Kristensen, R. M. & Møbjerg, N. (2009b).
Cyclomorphosis in Tardigrada: Adaption to
environmental constraints. Journal of Experimental
Biology, 212:2803-2811.
Halberg, K. A. & Møbjerg, N. (2012). First evidence of
epithelial transport in tardigrades: Comparative
investigation of organic anion transport. J. Exp. Biol.,
215:497-507.
Halberg K. A., Persson, D., Jørgensen, A. Kristensen,
R. M. and Møbjerg, N. submitted. Population
dynamics of a marine tardigrade: Temperature limits
geographical distribution of Halobiotus crispae.
Mar. Biol. Res.
Hengherr, S., Heyer, A. G., Köhler, H. R. & Schill, R.
(2009). Trehalose and anhydrobiosis in tardigrades –
evidence for divergence in response to dehydration.
FEBS. J., 275:281-288.
Hincha, D. K. & Hagemann, M. (2004). Stabilization
of model membranes during drying by compatible
solutes involved in the stress tolerance of plants and
microorganisms. Biochem. J. 383:277-283.
Hobsen, A. D., Stephenson, W. and Eden A. (1952).
Studies on the physiology of Ascaris lumbricoides:
II. The inorganic composition of the body fluid in
relation to that of the environment. J. Exp. Biol.,
29:22-29.
Hronowski, L. & Armstrong, J. B. (1977). Ion
composition of the plasma of Ambystoma
mexicanum. Comp. Biochem. Physiol. A, 58:181-183.
Jönsson, K. I. & Persson, O. (2010). Trehalose in three
species of desiccation tolerant tardigrades. Open
Zool. J., 3:1-5.
Jørgensen, A., Faurby, S., Hansen, J. G., Møbjerg, N.
and Kristensen, R. M. (2010). Molecular phylogeny
of Arthrotardigrada (Tardigrada). Mol. Phylogen.
Evol., 54:1006-1015.
Normant, M., Kubicka, M., Lapucki, T., Czarnowski,
W. and Michalowska M. (2005). Osmotic and ionic
haemolymph concentration in the Baltic Sea
amphipod Gammarus oceanicus in relation to water
salinity. Comp. Biochem. Physiol. A, 141:94-99.
Macallum, A. B. (1910). The inorganic composition of
the blood in vertebrates and invertebrates, and its
origin. Proc. R. Soc. Lond. B, 82(559):602-624.
Møbjerg N, Dahl C. (1996). Studies on the morphology
and ultrastructure of the Malpighian tubules of
Halobiotus crispae Kristensen, 1982 (Eutardigrada).
Zool. J. Linn. Soc., 116:85-99
Møbjerg, N., A. Jørgensen, J. Eibye-Jacobsen, K. A.
Halberg, D. Persson & R. M. Kristensen (2007).
New Records on cyclomorphosis in the marine
eutardigrade Halobiotus crispae (Eutardigrada:
Hypsibiidae). J. Limnol., 66 (suppl. 1): 132-140.
Møbjerg, N. M., Halberg, K. A., Persson., D.,
Jørgensen, A. & Kristensen R. M. (2011). Survival
in extreme environments – on current knowledge of
adaptations in tardigrades. Acta Physiol., 202: 409-
420.
Persson, D., Halberg K. A., Jørgensen A., Ricci C.,
Møbjerg N. & Kristensen R. M. (2010). Extreme
stress tolerance in tardigrades: Surviving space

conditions in low earth orbit. J. Zool. Syst. Evol.
Res., 49: 90-97.
Persson, D., Halberg K. A., Jørgensen A., Møbjerg
N. & Kristensen R. M. (2012). Neuroanatomy of
Halobiotus crispae: Tardigrade brain structure
suggests inclusion into Panarthropoda. J. Morphol.
(in review).
Robertson, J. D. (1949). Ionic regulation in some
marine invertebrates. J. Exp. Biol., 26:182-200.
Robertson, J. D. (1954). The chemical composition of
the blood of some aquatic chordates, including
members of the Tunicata, Cyclostomata and
Osteichthyes. J. Exp. Biol., 31:424-442.
Sands, M, Nicol, S. & McMinn, A. (1998). Fluoride in
Antarctic marine crustaceans. Mar. Biol., 132:591-
598.
Smith R. I. L. (1978). Summer and winter
concentrations of sodium, potassium and calcium in
some maritime Antarctic cryptogams. J. Ecol., 66
(3):891-909.
Sutcliffe, D. W. (1962). The composition of
haemolymph in aquatic insects. J. Exp. Biol.,
39:325-343.
Sutcliffe, D. W. (1963). The chemical composition of
haemolymph in insects and some other arthropods,
in relation to their phylogeny. Comp. Biochem.
Physiol., 9(2):121-135.
Tentori E. and Lockwood, A. P. M. (1990).
Haemolymph magnesium levels in some oceanic
crustaceans. Comp. Biochem. Physiol. A, 4:545-548.
Wilder, M. N., Ikuta, K., Atmomarsono M., Hatta,
T. and Komuro, K. (1998). Changes in osmotic
and ionic concentrations in the hemolymph of
Macrobrachium rosenbergii exposed to varying
salinities and correlation to ionic and crystalline
composition of the cuticle. Comp. Biochem. Physiol.
A, 119:941-950.
Wyatt, G. R. (1961). The biochemistry of insect
hemolymph. Ann. Rev. Entomol., 6:75-102.
Westh, P. and Kristensen, R. M. (1992). Ice formation
in the freeze-tolerant eutardigrades Adorybiotus
coronifer and Amphibolus nebulosus studied by
differential scanning calorimetry. Polar Biol.
12:693-699.
Westh, P. and Ramløv, H. (1991). Trehalose
accumulation in the tardigrade Adorybiotus
coronifer during anhydrobiosis. J. Exp. Zool.
258:303-311.
Yancey, P. H. (2005). Organic osmolytes as
compatible, metabolic and counteracting
cytoprotectants in high osmolarity and other
stresses. J. Exp. Biol., 208:2819-2830.
Inorganic ion composition in Tardigrada

Paper II

497
The Journal of Experimental Biology 215, 497-507
© 2012. Published by The Company of Biologists Ltd
doi:10.1242/jeb.065987
RESEARCH ARTICLE
First evidence of epithelial transport in tardigrades: a comparative investigation of
organic anion transport
Kenneth Agerlin Halberg* and Nadja Møbjerg
Department of Biology, The August Krogh Building, University of Copenhagen, Universitetsparken 13, DK-2100 Copenhagen Ø,
Denmark
*Author for correspondence (kahalberg@bio.ku.dk)
Accepted 31 October 2011
SUMMARY
We investigated transport of the organic anion Chlorophenol Red (CPR) in the tardigrade Halobiotus crispae using a new method for
quantifying non-fluorescent dyes. We compared the results acquired from the tardigrade with CPR transport data obtained from
Malpighian tubules of the desert locust Schistocerca gregaria. CPR accumulated in the midgut lumen of H. crispae, indicating that
organic anion transport takes place here. Our results show that CPR transport is inhibited by the mitochondrial un-coupler DNP
(1mmoll –1 ; 81% reduction), the Na + /K + -ATPase inhibitor ouabain (10mmoll –1 ; 21% reduction) and the vacuolar H + -ATPase inhibitor
bafilomycin (5mmoll –1 ; 21% reduction), and by the organic anions PAH (10mmoll –1 ; 44% reduction) and probenecid (10mmoll –1 ; 61%
reduction, concentration-dependent inhibition). Transport by locust Malpighian tubules exhibits a similar pharmacological profile,
albeit with markedly higher concentrations of CPR being reached in S. gregaria. Immunolocalization of the Na + /K + -ATPase -subunit
in S. gregaria revealed that this transporter is abundantly expressed and localized to the basal cell membranes. Immunolocalization
data could not be obtained from H. crispae. Our results indicate that organic anion secretion by the tardigrade midgut is transporter
mediated with likely candidates for the basolateral entry step being members of the Oat and/or Oatp transporter families. From our
results, we cautiously suggest that apical H + and possibly basal Na + /K + pumps provide the driving force for the transport; the exact
coupling between electrochemical gradients generated by the pumps and transport of ions, as well as the nature of the apical exit
step, are unknown. This study is, to our knowledge, the first to show active epithelial transport in tardigrades.
Key words: organic anion transport, Chlorophenol Red, 2,4-dinitrophenol, ouabain, bafilomycin, probenecid, para-aminohippuric acid, tardigrade,
insect, V-type H + -ATPase, Na + /K + -ATPase, Malpighian tubule.
INTRODUCTION
The ability to excrete metabolic waste products as well as
environmental toxins (xenobiotics) is a fundamental prerequisite
for animal life. One of the earliest identified systems involved in
such an excretion is the classic organic anion transport system
(Marshall and Vickers, 1923). In vertebrates, this system is
(especially) well known in the proximal tubule where it rids the
organism of various xenobiotic and endobiotic compounds,
through an ATP-dependent, net transepithelial secretory pathway
(reviewed by Dantzler, 2002; Burckardt and Burckardt, 2003;
Russel, 2010). The importance and understanding of similar
transport activities in various invertebrate phyla, however, is
extremely limited. In this study, we expand on the current
knowledge by investigating organic anion transport in the
tardigrade Halobiotus crispae (Tardigrada), as compared with the
desert locust Schistocerca gregaria (Arthropoda).
The phylum Tardigrada consists of a group of minute, eight-legged,
multicellular animals that, like arthropods and nematodes, belong to
the invertebrate superclade Ecdyzosoa (Aguinaldo et al., 1997). They
are considered essential to our understanding of early metazoan
evolution, yet fundamental questions concerning their basic biology
remain unanswered (Møbjerg et al., 2011). In spite of their small size
(~50–1200mm), tardigrades are relatively complex animals; they are
composed of >1000 cells, and possess a well-developed musculature
and nervous system, as well as a complex alimentary canal and
specialized reproductive and excretory organs (Dewel and Dewel,
1979; Rebecchi and Bertolani, 1994; Møbjerg and Dahl, 1996;
Schmidt-Raesa and Kulessa, 2007; Pelzer et al., 2007; Zantke et al.,
2008; Halberg et al., 2009a; Møbjerg et al., 2011; Rost-Roszkowska
et al., 2011). The alimentary canal can be divided into several
morphologically distinct regions i.e. bucco-pharyngeal apparatus,
oesophagus, midgut and hindgut, with the Malpighian tubules (MTs)
of eutardigrades positioned at the junction between these last two
sections. Interestingly, the same basic organizational pattern is found
in insects, and has been used as a strong argument for the homology
of these two organ systems (Greven, 1982; Møbjerg and Dahl, 1996).
Among multi-cellular animals, tardigrades exhibit an extraordinary
ability to resist environmental extremes, and are known to survive
conditions greatly exceeding those encountered in their natural habitat
– even in space (Jönsson et al., 2008; Rebecchi et al., 2008; Persson
et al., 2011). The biochemical and physiological mechanisms
mediating this unique tolerance, however, remain largely unidentified.
Previously, we have shown that the marine eutardigrade H. crispae
tolerates large changes in external salinity surviving periods of osmotic
stress by maintaining haemolymph osmotic pressure above that of
the external medium (Halberg et al., 2009b); an adaptive mechanism
likely present in all eutardigrades (Møbjerg et al., 2011). Here, we
identify organs involved in transepithelial transport of organic anions
and investigate the transport characteristics with the aim of providing
a better understanding of the unique biology in these animals.
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K. A. Halberg and N. Møbjerg
Organic anion transport has previously been described in other
groups of invertebrates (George et al., 1999; Torrie et al., 2004;
Faria et al., 2010). Notably, the alimentary canal and MTs of insects,
which collectively form the functional analogue of the vertebrate
kidney, have been studied (reviewed by Phillips, 1981; O’Donnell
et al., 2003; Dow and Davies, 2006). As in vertebrates, the excretory
organs of insects transport a wide range of organic solutes and
exogenous toxins through organic anion transporters (Oats), organic
anion-transporting peptides (Oatps), P-glycoproteins (Mdr/P-gp) as
well as multidrug resistance-associated proteins (Mrps) (Maddrell
et al., 1974; Bresler et al., 1990; Lanning et al., 1996; Linton and
O’Donnell, 2000; Torrie et al., 2004; Neufeld et al., 2005; Leader
and O’Donnell, 2005; O’Donnell and Leader, 2006; Chahine and
O’Donnell, 2009; Chahine and O’Donnell, 2010). Organic anions
are divided into type I and type II organic anions, on the basis of
structural and chemical properties (e.g. Wright and Dantzler, 2004).
The different groups of transporters vary in their transport
mechanisms, but overlap in their substrate specificity, as they
transport carboxylates and sulphonates interchangeably (Neufeld et
al., 2005; Chahine and O’Donnell, 2009). Oats and Oatps (solute
carriers belonging to the SLC22 and SLC21/SLCO family) transport
both small (450Da) hydrophobic type II organic anions (Oatps),
whereas Mdr/P-gp and Mrps (ABC transporters, ABCB and ABCC
subfamilies) generally transport large (>500Da) polyvalent type II
organic anions (Wright and Dantzler, 2004; Russel, 2010). From
the insect’s perspective, the clearance of exogenous toxins is of
particular interest, as insects often live in environments with high
xenobiotic exposure, potentially at harmful or lethal concentrations.
Accordingly, they are forced to process naturally occurring plant
toxins (Torrie et al., 2004; Neufeld et al., 2005), as well as
anthropogenic contaminants, such as insecticides (Lanning et al.,
1996; Neufeld et al., 2005; Buss and Callaghan, 2008). The
physiological importance of this detoxification system – and
implicitly the MTs – is emphasized by the fact that transcripts for
these transporters are enriched in the transcriptome of the MTs
(Wang et al., 2004), and because the expression of several Oatp and
Mdr transporters is significantly upregulated upon dietary exposure
to Oatp and Mdr substrates (Mulenga et al., 2008; Chahine and
O’Donnell, 2009; Chahine and O’Donnell, 2010). Indeed, it has been
suggested that organic solute excretion is the most significant
function of the insect MT (Dow and Davies, 2006). Nevertheless,
in spite of significant efforts in the past decades, the mechanisms
underlying transepithelial transport of organic anions is far from
understood, and our knowledge is limited to relatively few taxa.
In this study we examined epithelial transport in tardigrades.
Using a comparative approach, we investigated the sites,
characteristics and pharmacological profile of the net transepithelial
transport of Chlorophenol Red (CPR; 3,3-dichlorophenolsulphone-phthalein),
a pH indicator and a prototypic substrate of
the classic organic anion secretory pathway, in the tardigrade H.
crispae and the desert locust S. gregaria. Our results show that the
tardigrade midgut is the principal site of CPR transport, and that
this transport is active and transporter mediated. Additionally, our
data show that the pharmacological profiles of CPR transport in the
tardigrade midgut and locust MT are surprisingly similar.
MATERIALS AND METHODS
Quantification of CPR accumulation
The quantification of CPR accumulation was performed by
introducing a new method for quantifying non-fluorescent dyes. The
method exploits the optical properties of CPR by relating relative
Difference in red spectrum/green spectrum
relationship (%)
6000
5000
4000
3000
2000
1000
0
pH 6
r 2 =0.994
pH 7–10
r 2 =0.987
0 1 2 3 4 5
[CPR] (mmol l –1 )
pH 11
r 2 =0.996
Fig.1. Standard curves showing the percentage difference in the red
spectrum/green spectrum relationship as a function of Chlorophenol Red
(CPR) concentration at different pH values (pH6, 7–10 and 11). The curves
were fitted by regression analysis using OriginPro 7.5 with a third order
polynomial providing the best fit with r 2 values close to unity.
changes in spectral light to that of dye concentration (Fig.1). As
CPR exhibits a pH-dependent shift in colour (from yellow to violet)
between pH4.6 and 7.0, and a secondary shift (to purple) at pH>10,
a standard curve was constructed at three discrete colours relevant
to our investigation (red, violet and purple corresponding to
pH6–11). This was done in order to correct for potential pH
dependent effects on CPR quantification caused by the tissue. The
standard curves were constructed from optical analysis of
micrographs taken of samples (40ml) with known dye concentrations
(ranging from 0.1 to 5mmoll –1 ) at each respective pH value. The
optical measurements provided relative, arbitrary values for the light
intensities of the red, green and blue spectrum of spectral light from
each sample, and were acquired using the image analysis and
visualization software Imaris 6.4 (Bitplane, Zurich, Switzerland).
The standard curves are expressed as percentage difference in the
red spectrum/green spectrum relationship as a function of CPR
concentration (Fig.1). The curves were fitted by regression analysis
using OriginPro 7.5 (OriginLab, Northampton, MA, USA) with a
third order polynomial providing the best fit with r 2 values close to
unity. The CPR concentration in a given tissue was subsequently
calculated by employing the appropriate standard curve. The
standard curve was chosen by visually comparing the colour of the
accumulated CPR with that of the respective standard curves.
Contrary to our expectations, the midgut of H. crispae was the
only organ in which CPR accumulation was clearly visualized.
No accumulation was observed in the tardigrade MTs. As such,
quantification of dye accumulation was only performed from the
tardigrade midgut, and compared with the accumulation in the
MTs of S. gregaria. Specifically, 6–12 regions from areas of the
tardigrade midgut containing the highest dye intensity were
selected arbitrarily, in addition to a similar number from the
adjacent haemolymph, while four different regions were selected
from each insect MT. Each region selected represented a circle
with a diameter of 5mm for H. crispae and 25mm for S. gregaria.
The average light intensity within these circles was measured by
the Imaris program. Averaging the measured intensity of all
selected regions provided an overall average intensity for the red,
green and blue colour spectrum within each investigated organ
following each exposure. As CPR predominantly appears red (to
violet) in the midgut of H. crispae and in the MTs of S. gregaria,
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Epithelial transport in tardigrades
499
the red spectrum/green spectrum relationship offered an estimate
of dye accumulation, as calculated from the appropriate standard
curve. The final CPR concentration was normalized according to
the background light intensity, i.e. the tardigrade haemolymph or
untreated (control) insect MTs. For tardigrades, only regions
without gut content were selected for quantification, in order to
avoid the influence of gut content on the wavelength of captured
light. Deviations in animal depth did not influence CPR
quantification within the variation encountered in this study. The
MTs of S. gregaria consist of three structurally distinct regions,
i.e. proximal, middle and distal relative to the gut (Garret et al.,
1988). Dye accumulation was estimated from the proximal region
and parts of the middle region within 5mm of the junction with
the gut, as dye accumulation was highest here.
Test solutions
The haemolymph osmolality of H. crispae, kept at a salinity of
20p.p.t., was previously measured by nanolitre osmometry to
~950mOsmkg –1 (Halberg et al., 2009b). At present we do not know
the composition of tardigrade extracellular fluids. As such, the
experimental solution (control solution) was prepared from
evaporative reduction of seawater (SW; salinity 20p.p.t., pH8)
collected at the locality, and 35mmoll –1 glucose was added to
alleviate potential variation in experiments caused by differences
in nutrient availability. This yielded a final measured osmolality of
950±3mOsmkg –1 (N3). For S. gregaria an insect saline (control
solution) was prepared containing (in mmoll –1 ): 130 NaCl, 10 KCl,
4 NaHCO 3 , 2 MgSO 4 , 2 CaCl 2 , 6 NaH 2 PO 4 , 35 glucose and 5 Hepes,
titrated to pH7.2, with a measured osmolality of 336±2mOsmkg –1
(N3).
In order to explore the kinetics of CPR transport by MTs of S.
gregaria, a concentration–response curve was constructed over a
5000-fold range of CPR concentrations (1mmoll –1 to 5mmoll –1 )
(Fig.2). These data revealed that CPR transport is saturated at an
external CPR concentration of ~1.6mmoll –1 (V max 1.58mmoll –1 ,
K m 81.8mmoll –1 ; Fig.2). However, the MTs are unable to
concentrate the dye at high concentrations of CPR (>1mmoll –1 ;
Fig.2, inset). Experiments, on both animals, were performed at a
concentration of 1mmoll –1 CPR.
Test solutions were prepared from the two control solutions
containing 1mmoll –1 CPR, and one of the following inhibitors: 2,4-
dinitrophenol (DNP, 1mmoll –1 ), ouabain (10mmoll –1 ), bafilomycin
A 1 (5mmoll –1 ), para-aminohippuric acid (PAH; 10mmoll –1 ) or
probenecid (0.1–10mmoll –1 ). The final osmolality of the solutions
was measured on a Vapro 5520 vapour pressure osmometer (Wescor,
Logan, UT, USA). All solutions were titrated with NaOH to pH8
for H. crispae and pH7.2 for S. gregaria.
Experimental animals
Specimens of H. crispae Kristensen 1982 were collected on 11
February 2008 and 17 January 2010 at Vellerup Vig, Isefjord,
Denmark (55°44.206N, 11°51.258E) (see Fig.3A). Bottom samples
were collected with a mini van Veen grab at a depth of 1–2m (salinity
~20p.p.t., pH8). Rocks, algae and sediment collected with the grab
were freshwater shocked. The debris was decanted into a conical net
(mesh size 63mm) and subsequently transferred to SW from the
locality and kept at 4°C. Animals in the active stage (see Kristensen,
1982; Møbjerg et al., 2007; Halberg et al., 2009b) were identified
using a dissection microscope. The tardigrades were kept for a period
of up to 6months at 4°C in SW (salinity 20p.p.t., pH8) and regularly
supplied with fresh substrate, consisting mainly of sediment, organic
debris, filamentous algae and diatoms.
A
B
[CPR]lumen (mmol l –1 )
2.2
2.0
1.8
1.6
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0
tc
1 μmol l –1 10 μmol l –1 100 μmol l –1 1 mmol l –1 5 mmol l –1
lu
tc
lu
K m =81.8 μmol l –1
V max =1.58 mmol l –1
Lumen/bath ratio
tc
200
150
100
Specimens of S. gregaria Forskål 1775 were acquired from a
specialized animal shop (www.exopark.dk), and kept at room
temperature (RT) with light:dark periods of 16h:8h and fed annual
meadow grass (Poa annua) ad libitum.
Exposure to test solutions
Halobiotus crispae
Evaluation of CPR transport by tardigrade epithelia was performed
on whole animals immersed in the dye solution. Data obtained from
82 animals were used for the study – no distinction was made
between male and female specimens. Initial observations revealed
that the animals did not take up dye through the mouth or cuticle
(CPR non-punctured, Fig.3) and test solutions were therefore
introduced to the haemolymph of single specimens (length
300–500mm) through a small hole made in the cuticle in the anterior
part of the animal. The animal was incubated for a period of 60min
at RT in the respective test solution, and quickly washed in SW
prior to photography. Light micrographs of the specimens were taken
in bright-field at a 40 magnification, using an Olympus DP20
camera mounted on an Olympus BX50 microscope (Olympus,
Hamburg, Germany). Additionally, in order to investigate whether
the test solutions were ingested during the experimental period, intact
non-punctured animals were incubated in the CPR test solution for
a corresponding period. The animals were washed and photographed
as described above.
50
lu
tc
[CPR] bath (mmol l –1 )
lu
0
0 0.5 1
0 1 2 3 4 5
Fig.2. Accumulation of CPR as a function of external CPR concentration in
Malpighian tubules (MTs) of Schistocerca gregaria. (A)Representative light
micrographs of the MTs following exposure to different concentrations
(1mmoll –1 , 10mmoll –1 , 100mmoll –1 , 1mmoll –1 and 5mmoll –1 ) of CPR.
Scale bars, 100mm. lu, lumen; tc, trachea. (B)Luminal CPR concentration
as a function of bath CPR concentration, revealing the kinetic parameters
K m and V max for CPR transport. Each point shows the mean ± s.d. for
N4–6 animals with 3–5 MTs providing the estimate for each animal. The
solid line represents the fit to the Michaelis–Menten equation by non-linear
regression analysis (using error as weight). Insert shows the lumen/bath
ratio of CPR as a function of external CPR concentration.
lu
5
THE JOURNAL OF EXPERIMENTAL BIOLOGY

500
K. A. Halberg and N. Møbjerg
B
[CPR]luminal (mmol l –1 )
1.6
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0
SW
control
mg
C
(12)
SW control
CPR nonpunctured
mg
(6)
CPR non-punctured
mg
CPR
(13)
1 mmol l –1 CPR
+DNP
–81%
mg
**
(5)
+1 mmol l –1 DNP
+Bafilomycin
+Ouabain +DMSO
–21% –21%
n n
mg
*
+10 mmol l –1 ouabain
mg
*
(6) (8)
+5 μmol l –1 bafilomycin
A
+PAH
–44%
mg
*
*
**
(11)
+10 mmol l –1 PAH
mg
*
Leg 4
+Probenecid
–61%
mg
Leg 1
Leg 2
Leg 3
**
(10)
+10 mmol l –1 probenecid
Fig.3. CPR accumulation in the gut lumen of Halobiotus crispae,
and the effect of inhibitors on dye accumulation. (A)Light
micrograph of H. crispae from a ventral view showing the animalʼs
basic morphology. mg, midgut. Asterisks indicate MTs. Scale bar,
100mm. (B)Representative light micrographs of the midgut (mg)
following exposure to the various test solutions. Seawater control
shows the midgut of a punctured animal following immersion in
seawater, while CPR non-punctured shows the midgut of an intact
animal after immersion in CPR solutions. Estimations of luminal
CPR concentration were performed solely from areas devoid of gut
content, i.e. brown material in the gut. The percentage change in
CPR concentration, compared with experiments on CPR alone, is
noted. Scale bars, 50mm. (C)Corresponding luminal concentrations
of CPR. Data are depicted as means ± s.d. Asterisks refer to a
significant difference from CPR alone (*P

Epithelial transport in tardigrades
501
Microsystems, Wetzlar, Germany) with glass knives and
subsequently stained with Toluidine Blue.
For immunocytochemistry, whole animals of H. crispae and MTs
of S. gregaria were fixed in 3% paraformaldehyde in 0.1moll –1
sodium cacodylate buffer (pH7.4) for 60min and subsequently
transferred to 0.1moll –1 sodium cacodylate buffer. The tissue was
then dehydrated through a graded series of ethanol and xylene,
embedded in paraffin and sectioned into ~10mm sections, or
transferred to PBS and used as whole mounts for immunostaining.
Paraffin sections were deparaffinized through a graded series of xylene
and alcohol, washed in saline (control solution; see ‘Test solutions’
above) and blocked with 10% normal goat serum (Invitrogen,
Carlsbad, CA, USA) for 30min, prior to incubation with primary
antibody. Paraffin sections, as well as whole mounts, were incubated
overnight at 4°C in insect saline (MTs) or PBS (tardigrades) containing
10% normal goat serum, 0.1% Triton-X and primary antibody. The
Na + /K + -ATPase -subunit monoclonal mouse primary antibody 5-
IgG (10mgml –1 ) was developed by D. M. Famborough, and obtained
from the Developmental Studies Hybridoma Bank (University of
Iowa, Iowa City, IA, USA). This antibody has been used to identify
the Na + /K + -ATPase in numerous excretory tissues, including MTs of
Drosophila melanogaster (Lebovitz et al., 1989; Torrie et al., 2004),
the gills of the blue crab Callinectes sapidus (Towle et al., 2001) and
the pronephros of Ambystoma mexicanum (Haugen et al., 2010).
Following an extensive wash in saline, the tissue was incubated with
(anti-mouse) Alexa Fluor 594 (1:100) secondary antibody (Invitrogen)
overnight at 4°C. The tissue was rinsed with saline then counterstained
with Alexa Fluor 488-conjugated phalloidin (luminal marker; 1:40;
Invitrogen) and DAPI (50mgml –1 ; Invitrogen) for 2h, washed and
mounted on glass coverslips in Vectashield (Vector Laboratories Inc.,
Burlingame, CA, USA). Images were acquired using a Leica DM
RXE 6 TL microscope equipped with a Leica TCS SP2 AOBS
confocal laser scanning unit (Leica Microsystems, Wetzlar, Germany).
The tissue was scanned employing sequential scanning (setting:
between frames) using the 488nm line of an argon/krypton laser and
the 594nm line of a helium laser, in addition to the 405nm UV laser
line. The image series was processed and edited using Imaris software.
Confocal images are based on 240 optical sections of a Z-series
performed at intervals of 0.7mm. Experiments were conducted
multiple times with corresponding results. All control preparations
without primary antibody were negative for immunostaining.
Chemicals
All chemicals were obtained from Sigma-Aldrich (St Louis, MO,
USA). Bafilomycin was dissolved and stored in dimethyl sulphoxide
(DMSO). Inhibitors were allowed to dissolve in the CPR solution
overnight prior to use.
Statistics
Data are expressed as means ± s.d. unless otherwise stated. The
statistical significance of differences between the various exposures
was tested using one-way ANOVA followed by a Tukey multiple
comparisons of means. The statistical tests were performed using
the data analysis program OriginPro 7.5 (OriginLab). Significance
levels were P>0.05 (not significant), P

502
K. A. Halberg and N. Møbjerg
[CPR]lumen (mmol l –1 )
A
1.6
1.4
1.2
1.0
0.8
0.6
0.4
CPR
mg
B
(13)
*
+Bafilomycin
+DMSO
–14%
+DMSO
–21%
CPR
mg
**
*
gm
(5) (8)
C
tc lu lu tc lu
1.6
1.4
1.2
1.0
0.8
0.6
0.4
D
+DMSO
+5%
**
(11) (10)
+Bafilomycin
+DMSO
–26%
**
(8)
tc
Fig.6. The effect of bafilomycin and dimethylsulphoxide (DMSO) on
CPR accumulation. (A)Representative light micrographs of the
tardigrade midgut (mg) following exposure to the test solutions.
The percentage change in CPR concentration, compared with
experiments with CPR alone, is noted. Scale bars, 50mm.
(B)Corresponding luminal concentration of CPR in midgut lumen.
(C)Representative light micrographs of locust MTs following
exposure to the test solutions. The percentage change in CPR
concentration, compared with experiments on CPR alone, is noted.
Scale bars, 100mm. (D)Corresponding luminal concentration of
CPR in the MTs. Asterisks refer to significant difference from
1mmoll –1 CPR alone (*P

Epithelial transport in tardigrades
503
A
[CPR]lumen (mmol l –1 )
1.6
1.4
1.2
1.0
0.8
0.6
0.4
0.2
+0.1 mmol l –1
CPR –26%
CPR
mg
B
(13)
**
mg
** **
(7)
+1 mmol l –1
–56%
*
mg
**
+10 mmol l –1
–61%
mg
(4) (10)
C
tc
lu
lu
1.6 D ** **
(11) **
1.4
1.2
1.0
0.8
0.6
0.4
0.2
CPR
+0.1 mmol l –1
–9%
(5)
+1 mmol l –1
–47%
lu
(5)
+10 mmol l –1
–77%
tc
**
lu
(9)
Fig.7. Titration of the inhibitory effect of
probenecid. (A)Representative light
micrographs of the tardigrade midgut (mg)
following exposure to the different
concentrations (0.1, 1 and 10mmoll –1 ) of
probenecid. The percentage change in CPR
concentration, compared with experiments
with CPR alone, is noted. Scale bars, 50mm.
(B)Corresponding luminal concentration of
CPR in midgut lumen. (C)Representative
light micrographs of the insect MTs following
exposure to the different concentrations (0.1,
1 and 10mmoll –1 ) of probenecid. The
percentage change in CPR concentration,
compared with experiments on CPR alone, is
noted. Scale bars, 100mm. (D)Corresponding
luminal concentration of CPR in the MTs.
Asterisks refer to significant difference from
1mmoll –1 CPR alone (*P

504
K. A. Halberg and N. Møbjerg
probenecid concentration and CPR accumulation (and activity; data
not shown). At a concentration of 0.1mmoll –1 probenecid, CPR
concentration was reduced by 26% with a luminal concentration of
0.42±0.07mmoll –1 in H. crispae, and by 9% corresponding to a
concentration of 1.2±0.14mmoll –1 in S. gregaria. When applied at
a concentration of 1mmoll –1 , probenecid reduced CPR concentration
by 56% corresponding to a luminal concentration of
0.25±0.05mmoll –1 in H. crispae, and by 47% equivalent to
0.7±0.16mmoll –1 in S. gregaria (Fig.7).
Midgut and MT structure and immunostaining
Our transport studies indicate that CPR transport is active and
transporter mediated in the midgut of H. crispae as well as the MTs
of S. gregaria. It is well established that the vacuolar H + -ATPase
is important for energizing transepithelial transport in insects,
whereas (and in contrast to vertebrate literature) the role of the
Na + /K + -ATPase is controversial (e.g. Torrie et al., 2004; Beyenbach
et al., 2010). The data obtained in the present study using ouabain
would suggest that the Na + /K + -ATPase is important for CPR
transport in both animals. Conversely, ouabain could act as a
competitive inhibitor, being transported by the same transporters as
CPR. We therefore investigated whether the Na + /K + pump is
expressed in the epithelia.
Unfortunately, our attempt to localize Na + /K + -ATPase in
tardigrades was unsuccessful, both on whole mounts and on paraffin
sections, and also in attempts at antigen retrieval.
Our observations on the MTs of S. gregaria generally correspond
with those previously described (Garret et al., 1988), when focusing
on the proximal region and parts of the middle region used in the
present study. In general, we observed a prominent brushborder
(Fig.8B,C) and at least two cell types. One type, a principal-like cell,
has a single nucleus, while the other is a very large double-nucleated
cell (dashed circles in Fig.8). Cross-striated muscle fibers are present
in both the transverse (Fig.8A,B) and longitudinal direction
(Fig.8E,G). The transverse muscle fibers were only observed in the
proximal-most region of the MTs, whereas the longitudinal muscle
extended the entire length of the tubule. Visual observations revealed
that the transverse muscles aid the clearance of luminal content into
the midgut through peristaltic contractions. Our immunocytochemical
analysis shows that the Na + /K + -ATPase is expressed in the MTs of
S. gregaria, and is localized to the basal plasma membranes
(Fig.8D–G), although to a lesser extent in the double-nucleated cells
(Fig.8D,G). The localization was confirmed by counterstaining of
nuclei by DAPI and staining apical microvilli with phalloidin. This
is, to our knowledge, the first study showing immunolocalization of
the Na + /K + -ATPase in S. gregaria MTs. Interestingly, the localization
of the transporter substantiates previously published data on isolated
basal membrane fractions from MTs and hindgut from S. gregaria,
which show a prominent Na + /K + -ATPase activity (Al-Fifi, 2007).
DISCUSSION
Our study is significant in two main aspects: (i) we are the first to
provide evidence for active epithelial transport in tardigrades, and
(ii) our data show that tardigrades possess an organic anion transport
system. Consequently, our study is important for understanding the
evolution of transport systems.
CPR is categorized as a sulphonate, but is known to compete with
substrates for both solute carriers (SLC22 and SLC21/SLCO) and
ABC transporters in a number of organisms (Pritchard et al., 1999;
Linton and O’Donnell, 2000; Chahine and O’Donnell, 2009). In H.
crispae, CPR transport is ATP dependent (strong inhibition by DNP),
probably energized by both the Na + /K + -ATPase and the V-type H + -
ATPase (bafilomycin and ouabain reduce CPR accumulation), and
possibly transporter mediated (inhibited by the prototypic organic
anions PAH and probenecid). The latter suggests that CPR transport
is (at least partly) transcellular. Comparing the transport characteristics
of CPR transport between the tardigrade midgut and the insect MT
reveals a surprisingly similar overall pharmacological profile of the
investigated tissues, albeit with markedly higher concentrations of
CPR observed in the insect MT. In addition to transcellular transport
of the dye, fluid secretion may augment transport of organic anions
by convective secretion through the paracellular pathway, and/or
reduce diffusive back-flux of organic anions from the tubule lumen
to the haemolymph (O’Donnell and Leader, 2006; Chahine and
O’Donnell, 2010). Consequently, without a measure of fluid secretion
rates, and thereby a measure of net CPR secreted, we cannot conclude
whether the difference in relative concentration of CPR in the gut
lumen of H. crispae, compared with the MTs of S. gregaria, is a
consequence of a lower dye transport capacity, or whether it reflects
differences in fluid transport rates. Also, we cannot be certain that
the concentration of CPR in the haemolymph of H. crispae is exactly
the same as that of the surrounding bath – although, the relatively
high dye concentration used along with the small diffusion distances
involved, the relatively long exposure time (60min) and the animal
movements that facilitate fluid exchange, would make potential
concentration differences negligible.
A vast number of papers have investigated various aspects of
organic anion transport in insect MTs. In the present study we
observed a mean CPR concentration of 1.3±0.13mmoll –1 when no
inhibitors were added. This concentration is elevated above the bath
concentration (1mmoll –1 ), albeit by a much smaller factor than that
reported from other insects (Maddrell et al., 1974; Bresler et al.,
1990; Linton and O’Donnell, 2000; Leader and O’Donnell, 2005).
However, when exposed to external CPR concentrations of 1, 10
and 100mmoll –1 , the mean luminal CPR concentrations were
0.16±0.03, 0.73±0.09 and 0.89±0.07mmoll –1 , respectively, which
is a factor of ~160, 70 and 9 above bath concentrations (Fig.2).
When exposed to a bath concentration of 5mmoll –1 CPR, the mean
luminal concentration was 1.79±0.33mmoll –1 , a factor of 2.8 below
external concentrations. Consequently, the luminal CPR
concentration is maximally elevated (~160-fold) above that in the
bathing medium when the latter contains CPR at a concentration of
1mmoll –1 . This concentrative ability is among the highest measured
for insects, and is an additional confirmation of active transport of
CPR by the insect MTs.
Ouabain, a well-characterized, potent inhibitor of the Na + /K + -
ATPase, reduced the CPR concentration in both the tardigrade
midgut and the insect MTs by ~20–21%. A similar 23% reduction
in PAH secretion in the presence of 1mmoll –1 ouabain was reported
from MTs of D. melanogaster (Linton and O’Donnell, 2000), while
fluorescein uptake was reduced in MTs of Blaberus giganteus by
30% and in MTs of Locusta migratoria by 20%, when ouabain was
used at 1 and 0.1mmoll –1 , respectively (Bresler et al., 1990). Our
immunocytochemical investigation on the MTs of S. gregaria
revealed expression of the Na + /K + -ATPase in the basal cell
membranes, although to a lesser extent in the double-nucleated cells.
A basal localization of the Na + /K + -ATPase in principal cells of MTs
of D. melanogaster has previously been reported (Torrie et al., 2004).
The fact that there is expression of the Na + /K + -ATPase in the basal
membranes, and that CPR transport is ouabain sensitive, suggests
that the pump is important for CPR transport. Alternatively, it could
be argued that the inhibitory effects of ouabain are due to competitive
inhibition, rather than to non-competitive inhibition, given the fact
that ouabain is actively transported by members of the SLC21/SLCO
THE JOURNAL OF EXPERIMENTAL BIOLOGY

Epithelial transport in tardigrades
505
subfamily (Oatps) in D. melanogaster (Torrie et al., 2004). Whether
this is ubiquitous in insects is at present not known; however, several
members of the Oatp family are known to transport ouabain in
human tissue (see Hagenbuch and Gui, 2008). Competition studies
using fluorescently labeled ouabain would help in clarifying this
matter. Considering the striking similarities between the
pharmacological profiles of the tardigrade midgut and insect MT,
it seems reasonable to assume that the Na + /K + -ATPase is similarly
present in the basal cell membranes of the tardigrade midgut (in
spite of failed attempts to localize this transporter). In support of
this interpretation is the fact that transcripts for the -subunit of the
Na + /K + -ATPase were found in the expressed sequence tags (EST)
library from Hybsibius dujardini Doyère, 1840 (TardiBASE cluster
ID: HDC01733 TardiBASE; http://xyala.cap.ed.ac.uk/research/
tardigrades/tardibase.shtml).
Bafilomycin is a specific inhibitor of the V-type H + -ATPase and
was found to reduce CPR accumulation in both H. crispae and S.
gregaria. This observation is consistent with the fact that the V-type
H + -ATPase is viewed as being central to the transport activities of
the MT in insects (Weng et al., 2003; Beyenbach et al., 2010) and
perhaps also for transport in tardigrades; the B-, C-, D-, E-, G- and
H-subunits of the V-type H + -ATPase were found in the EST library
of H. dujardini (TardiBASE). Indeed, visual changes to CPR colour
were observed in the midgut of H. crispae (CPR became purple; data
not shown) 10min post-incubation, suggesting an alkalization of the
midgut content. This observation is in accordance with specific
inhibition of an apical V-type H + -ATPase in the tardigrade midgut.
PAH is a prototypical substrate of the classic organic anion transport
system (i.e. the SLC22 subfamily) and was shown to be an effective
inhibitor of CPR transport in both H. crispae and S. gregaria (~40%
reduction in both animals). This finding is a strong indication that
PAH (carboxylate) and CPR (sulphonate) transport are mediated by
a common transporter, or alternatively through two separate transport
systems overlapping in affinity, in both investigated epithelia. Whether
carboxylates (e.g. PAH and probenecid) and sulphonates (e.g. CPR)
are handled by a common transporter (Bresler et al., 1990), or by two
separate transport systems (Maddrell et al., 1974; Linton and
O’Donnell, 2000; Chahine and O’Donnell, 2009), appears to be highly
variable and/or species specific. Indeed, transport of PAH and
probenecid (carboxylates) and CPR and methotrexate (MTX;
sulphonates) are all mediated by one tranporter (OAT1) in humans,
but not in rats (Burckardt and Burckardt, 2003), whereas MTX uptake
is competitively inhibited by CPR and probenecid, but not PAH in
D. melongaster (Cahine and O’Donnell, 2009).
Probenecid is implicitly regarded as a competitive inhibitor of
organic anion transport, and has been shown to compete with both
solute carrier and ABC transporter substrates (Bresler et al., 1990;
Linton and O’Donnell, 2000; Neufeld et al., 2005; Leader and
O’Donnell, 2005). It is typically applied in a concentration of
1mmoll –1 ; here, we examined the effects of probenecid on CPR
transport in the range 0.1–10mmoll –1 . In H. crispae, probenecid
reduced CPR accumulation by 26%, 56% and 61% at a concentration
of 0.1, 1 and 10mmoll –1 , respectively, whereas the corresponding
reduction in CPR concentration was 9%, 47% and 77% in S.
gregaria. In the presence of very high probenecid concentrations
(10mmoll –1 ), we observed an almost total loss of motility in H.
crispae. Indeed, the level of animal movement was comparable to
that in treatments containing DNP. This observation suggests that
the drug affected processes in addition to CPR transport at the given
concentration. In fact, when applied at concentrations of 1mmoll –1
or above, probenecid is reported to induce a range of non-specific
effects presumably initiated by the uncoupling of mitochondrial
Haemolymph
Na + /K + -ATPase
K + OA –
ATP
P Na +
i +ADP
Tardigrade
midgut cell
P i +ADP H+ ATP
V-type H + -ATPase
Lumen
Solute carrier
(SLC21/SLCO)
Nucleus
OA –
Basal lamina
Apical junction
complex
Fig.9. Tentative model of the tardigrade midgut cell derived from the
current study on H. crispae. Based on the pharmacological profile of the
tardigrade midgut epithelium, both the Na + /K + -ATPase and the V-type H + -
ATPase are potential candidates for providing an electrochemical driving
force for the transepithelial movement of organic anions. Transport
characteristics and the presence in tardigrade EST libraries suggest that a
member of the SLC21/SLCO transporter family may mediate the
basolateral entry of organic anions in tardigrades. The exact coupling
between electrochemical gradients generated by the pumps and transport
of ions, as well as the nature of the apical exit step, are not known.
oxidative phosphorylation (Masereeuw et al., 2000). It is therefore
possible that the reduction in CPR transport is non-specific when
the drug is applied in concentrations ≥1mmoll –1 . At lower
concentrations of probenecid (

506
K. A. Halberg and N. Møbjerg
ID: HDC00004, HDC02687 and HDC03352) (ABCB and ABCC
protein families) are present and expressed in tardigrades – just as
they are in insects (Maddrell et al., 1974; Lanning et al., 1996;
Bresler et al., 1990; Linton and O’Donnell, 2000; Torrie et al., 2004;
Neufeld et al., 2005; Leader and O’Donnell, 2005; O’Donnell and
Leader, 2006; Chahine and O’Donnell, 2009). Comparing the
transport characteristics of the different groups of transporters reveals
that it is unlikely that members of the ABC transporter family (i.e.
P-gps and Mrps) mediate the basolateral entry of CPR in H. crispae
and S. gregaria. This assumption is based on the fact that P-gp
transporters predominantly recognize large cationic species (Russel,
2010), and because Mrp transporters mainly transport large
(>500Da) polyvalent type II organic anions (Wright and Dantzler,
2004; Russel, 2010) – the apical transporter MRP2, however, was
shown to transport PAH in humans (see below). In addition, PAH
does not compete with MTX (an Mrp substrate) in D. melongaster,
suggesting distinct transport systems for these compounds in insects
(Chahine and O’Donnell, 2009). In contrast, both members of the
SLC22 and SLC21/SLCO protein families (Oats and Oatps) have
been reported to transport all the investigated organic anions,
including probenecid, PAH and CPR (Pritchard et al., 1999; Lee
and Kim, 2004; Torrie et al., 2004). Therefore, it seems probable
that an Oat or Oatp homologue mediates the basolateral entry of
CPR in the tardigrade midgut cell as well as the insect MTs (Fig.9).
At present, our data do not allow us to make assumptions on the
nature of the luminal exit. Evidence suggests that MRP2 is involved
in the efflux of organic anions across the brush-border membrane
in the human kidney proximal tubule (Leier et al., 2000), and a
similar situation could exist in tardigrades and insects. Our data
tentatively suggest that the V-type H + -ATPase, and perhaps also
the Na + /K + -ATPase, provide the driving force for the transepithelial
transport of organic anions in both H. crispae and S. gregaria
(Fig.9). It is likely that a large lumen-positive transepithelial
potential, generated by an apical H + -ATPase provides a significant
driving force for the accumulation of anions in the lumen. However,
the exact coupling between electrochemical gradients generated by
the pumps and transport of the ions is not known.
In future studies, it would be of interest to investigate whether
substrates of Mrps (e.g. Texas Red and MTX) also accumulate in
the midgut of tardigrades, and whether transport of these anions
occurs via a separate or a common transporter to CPR. Similarly,
an understanding of the electrophysiological properties of the
midgut epithelium would be highly relevant in our ongoing struggle
to understand the complex biology of these amazing animals.
ACKNOWLEDGEMENTS
We would like to thank Reinhardt M. Kristensen for the use of the Olympus BX50
stereomicroscope, Jette Lyby Michelsen for technical assistance and Dennis K.
Persson and Aslak Jørgensen for help during sampling.
FUNDING
Funding came from the 2008 Faculty of Science, University of Copenhagen Freja-
Programme and from the Carlsberg Foundation.
REFERENCES
Aguinaldo, A. M. A., Turbeville, J. M., Linford, L. S., Rivera, M. C., Garey, J. R., Raff,
R. A. and Lake, J. A. (1997). Evidence for a clade of nematodes, arthropods and other
moulting animals. Nature 387, 489-493.
Al-Fifi, Z. I. A. (2007). Comparative of effect of inhibitors on the ATPase from the
excretory systems of the usherhopper, Poekilocerus bufonius and desert locust,
Schistocerca gregaria. Am. J. Cell Biol. 2, 11-22.
Beyenbach, K. W., Skaer, H. and Dow, J. A. T. (2010). The developmental, molecular,
and transport biology of Malpighian tubules. Annu. Rev. Entomol. 55, 351-374.
Bresler, V. M., Belyaeva, E. A. and Mozhayeva, M. G. (1990). A comparative study on
the system of active transport of organic acids in Malpighian tubules of insects. J. Insect
Physiol. 36, 259-270.
Burckardt, B. C. and Burckardt, G. (2003). Transport of organic anions across the
basolateral membrane of proximal tubule cells. Rev. Physiol. Biochem. Pharmacol. 146,
95-158.
Buss, D. S. and Callaghan, A. (2008). Interaction of pesticides with p-glycoprotein and
other ABC proteins: A survey of the possible importance to insecticides, herbicide and
fungicide resistance. Pestic. Biochem. Physiol. 90, 141-153.
Chahine, S. and OʼDonnell, M. J. (2009). Physiological and molecular characterization of
methotrexate transport by Malpighian tubules of adult Drosophila melanogaster. J. Insect
Physiol. 55, 927-935.
Chahine, S. and OʼDonnell, M. J. (2010). Effects of acute or chronic exposure to dietary
organic anions on secretion of methotrexate and salicylate by Malpighian tubules of
Drosophila melanogaster larvae. Arch. Insect Biochem. Physiol. 73, 128-147.
Dantzler, W. H. (2002). Renal organic anion transport: a comparative and cellular
perspective. Biochim. Biophys. Acta 1566, 169-181.
Dewel, R. A. and Dewel, W. C. (1979). Studies on the tardigrades. J. Morphol. 161, 79-
110.
Dow, J. A. T. and Davies, S. A. (2006). The Malpighian tubule: rapid insights from postgenomic
biology. J. Insect. Physiol. 52, 365-378.
Faria, M., Navarro, A., Luckenbach, T., Piña, B. and Barata, C. (2010). Characterization
of the multixenobiotic resistance (mxr) mechanism in embryos and larvae of the zebra
mussel (Dreissena polymorpha) and studies on its role in tolerance to single and mixture
combinations of toxicants. Aqua. Toxicol. 101, 78-87.
Garret, M. A., Bradley, T. J., Meredith, J. E. and Phillips, J. E. (1988). Ultrastructure of
the Malpighian tubules of Schistocerca gregaria. J. Morphol. 195, 313-325.
George, R. L., Wu, X., Huang, W., Fei, Y. J., Leibach, F. H. and Ganapathy, V. (1999).
Molecular cloning and functional characterization of a polyspecific organic anion
transporter from Caenorhabditis elegans. J. Pharmacol. Exp. Ther. 291, 596-603.
Greven, H. (1982). Homologues or analogues A survey of some structural patterns in
Tardigrada. In Proceedings of the Third International Symposium on the Tardigrada (ed.
D. R. Nelson), pp. 55-76. Johnson City, Tennessee: East Tennessee State University
Press.
Hagenbuch, B. and Gui, C. (2008). Xenobiotic transporters of human organic anion
transporting polypeptides (OATP) family. Xenobiotica 38, 778-801.
Halberg, K. A., Persson, D., Møbjerg, N., Wanninger, A. and Kristensen, R. M.
(2009a). Myoanatomy of the marine Tardigrade Halobiotus crispae (Eutardigrada:
Hypsibiidae). J. Morphol. 270, 996-1013.
Halberg, K. A., Persson, D., Ramløv, H., Westh, P., Kristensen, R. M. and Møbjerg, N.
(2009b). Cyclomorphosis in Tardigrada: adaption to environmental constraints. J. Exp.
Biol. 212, 2803-2811.
Haugen, B. M., Halberg, K. A., Jespersen, Å., Prehn, L. R. and Møbjerg, N. (2010).
Functional characterization of the vertebrate primary ureter: structure and ion transport
mechanisms of the pronephric duct of axolotl larvae (Amphibia). BMC Develop. Biol. 10,
56.
Jönsson, K. I., Rabbow, E., Schill, R. O., Ringdahl, M. H. and Rettberg, P. (2008).
Tardigrades survive exposure to space in low Earth orbit. Curr. Biol. 18, R729-R731.
Kristensen, R. M. (1982). The first record of cyclomorphosis in Tardigrada based on a
new genus and species from Arctic meiobenthos. Z. zool. Syst. Evolut.-forsch. 20, 249-
270.
Lanning, C. L., Fine, R. L., Corcoran, J. J., Ayad, H. M., Rose, R. L. and Abou-Donia,
M. B. (1996). Tobacco budworm P-glycoprotein: biochemical characterization and its
involvement in pesticide resistance. Biochim. Biophys. Acta. 1291, 155-162.
Leader, J. P. and OʼDonnell, M. J. (2005). Transepithelial transport of fluorescent p-
glycoprotein and MRP2 substrates by insect Malpighian tubules: confocal microscopic
analysis of secreted fluid droplets. J. Exp. Biol. 208, 4363-4376.
Lebovitz, R. M., Takeyasu, K. and Fambrough, D. M. (1989). Molecular characterization
and expression of the (Na + + K + )-ATPase alpha-subunit in Drosophila melanogaster.
EMBO J. 8, 193-202.
Lee, W. and Kim, R. B. (2004). Transport and renal drug elimination. Annu. Rev.
Pharmacol. Toxicol. 44, 137-166.
Leier, I., Hummel-Eisenbeiss, J., Cui, Y. and Keppler, D. (2000). ATP-dependent paraaminohippurate
transport by apical multidrug resistance protein MRP2. Kidney Int. 57,
1636-1642.
Linton, S. M. and OʼDonnell, M. J. (2000). Novel aspects of the transport of organic
anions by Malpighian tubules of Drosophila melanogaster. J. Exp. Biol. 203, 3575-3584.
Maddrell, S. H. P., Gardiner, B. O. C., Pilcher, D. E. M. and Reynolds, S. E. (1974).
Active transport by insect Malpighian tubules of acidic dyes and of acylamides. J. Exp.
Biol. 61, 357-377.
Marshall, E. K., Jr and Vickers, J. L. (1923). The mechanism of the elimination of
phenolsulphonephthalein by the kidney - a proof of secretion by the convoluted tubules.
Bull. Johns Hopkins Hosp. 34, 1-6.
Masereeuw, R., van-Pelt, A. P., van-Os, S. H. G., Willems, P. H. G. M., Smits, P. and
Russel, F. G. M. (2000). Probenecid interferes with renal oxidative metabolism: a
potential pitfall in its use as an inhibitor of drug transport. Br. J. Pharmacol. 131, 57-62.
Møbjerg, N. and Dahl, C. (1996). Studies on the morphology and ultrastructure of the
Malpighian tubules of Halobiotus crispae Kristensen 1982 (Eutardigrada). Zool. J. Linn.
Soc. 116, 85-99.
Møbjerg, N., Jørgensen, A., Eibye-Jacobsen, J., Halberg, K. A., Persson, D. and
Kristensen, R. M. (2007): New records on cyclomorphosis in the marine eutardigrade
Halobiotus crispae (Eutardigrada: Hypsibiidae). J. Limnol. 66 Suppl.1, 132-140.
Møbjerg, N. M., Halberg, K. A., Jørgensen, A., Persson. D., Bjørn, M., Ramløv, H. and
Kristensen, R. M. (2011). Survival in extreme environments – on the current knowledge
of adaptations in tardigrades. Acta Physiologica 202, 409-420.
Mulenga, A., Khumthong, R., Chalaire, K. C., Strey, O. and Teel, P. (2008). Molecular
and biological characterization of the Amblyomma americanum organic anion transporter
polypeptide. J. Exp. Biol. 211, 3401-3408.
Neufeld, D. S. G., Kaufmann, R. and Kurtz, Z. (2005). Specificity of the fluorescein
transport process in Malpighian tubules of the cricket Acheta domesticus. J. Exp. Biol.
208, 2227-2236.
OʼDonnell, M. J. and Leader, J. P. (2006). Changes in fluid secretion rate alter net
transepithelial transport of MRP2 and p-glycoprotein substrates in Malpighian tubules of
Drosophila melanogaster. Arch. Insect Biochem. Physiol. 63, 123-134.
THE JOURNAL OF EXPERIMENTAL BIOLOGY

Epithelial transport in tardigrades
507
OʼDonnell, M. J., Ianowski, J. P., Linton, S. T. and Rheault, M. R. (2003). Inorganic and
organic anion transport by insect renal epithelia. Biochim. Biophys. Acta 1618, 194-206.
Pelzer, B., Dastych, H. and Greven, H. (2007). The osmoregulatory/excretory organs of
the glacier-dwelling eutardigrade Hypsibius klebelsbergi Mihelcic, 1959 (Tardigrada).
Mitt. Hamb. Zool. Mus. Inst. 104, 61-72.
Persson, D., Halberg, K. A., Jørgensen, A., Ricci, C., Møbjerg, N. and Kristensen, R.
M. (2011). Extreme stress tolerance in tardigrades: surviving space conditions in low
earth orbit. J. Zool. Syst. Evol. Res. 49, 90-97.
Phillips, J. (1981). Comparative physiology of insect renal function. Am. J. Physiol. Regul.
Integr. Comp. Physiol, 241, R241-R257.
Pritchard, J. B., Sweet, D. H., Miller, D. S. and Walden, R. (1999). Mechanism of
organic anion transport across the apical membrane of choroid plexus. J. Biol. Chem.
274, 33382-33387.
Rebecchi, L. and Bertolani, R. (1994). Maturative pattern of ovary and testis in
eutardigrades of fresh-water and terrestrial habitats. Invert. Rep. Dev. 26, 107-117.
Rebecchi, L., Altiero, T., Guidetti, R., Cesari, M., Bertolani, R., Negroni, M. and
Rizzo, A. M. (2008). Tardigrade resistance to space effects: first results of
experiments on the LIFE-TARSE mission on FOTON-M3 (September 2007).
Astrobiology 6, 581-591.
Rost-Roszkowska, M. M., Poprawa, I., Wójtowicz, M. and Kaczmare, L. (2011).
Ultrastructural changes of the midgut epithelium in Isohypsibius granulifer granulifer
Thulin, 1928 (Tardigrada: Eutardigrada) during oogenesis. Protoplasma 248, 405-
414.
Russel, F. G. M. (2010). Transporters: importance in drug absorption, distribution, and
removal. In Enzyme- and Transporter-Based Drug-Drug Interactions (ed. K. S. Pang, A.
Rodrigues and R. M. Peter), pp. 27-49. Heidelberg: Springer.
Schmidt-Rhaesa, A. and Kulessa, J. (2007). Muscular architecture of Milnesium
tardigradum and Hypsibius sp. (Eutardigrada, Tardigrada) with some data on
Ramazzottius oberhaeuseri. Zoomorphology 126, 265-281.
Torrie, L. S., Radford, J. C., Southall, T. D., Kean, L., Dinsmore, A. J., Davies, S. A.
and Dow, J. A. T. (2004). Resolution of the insect ouabain paradox. Proc. Natl. Acad.
Sci. USA. 101, 13689-13693.
Towle, D. W., Paulsen, R. S., Weihrauch, D., Kordylewski, M., Salvador, C., Lignot, J.
H. and Spannings-Pierrrot, C. (2001). Na + +K + -ATPase in gills of the blue crab
Callinectes sapidus: cDNA sequencing and salinity-related expression of -subunit
mRNA and protein. J. Exp. Biol. 204, 4005-4012.
Wang, J., Kean, L., Yang, J., Allan, A. K., Davies, S. A., Herzyk, P. and Dow, J. A. T.
(2004). Function-informed transcriptome analysis of Drosophila renal tubule. Gen. Biol.
5, R69.
Weng, X.-H., Huss, M., Wieczorek, H. and Beyenbach, K. W. (2003). The V-type H + -
ATPase in Malpighian tubules of Aedes aegypti: localization and activity. J. Exp. Biol.
206, 2211-2219.
Wright, S. W. and Dantzler, W. H. (2004). Molecular and cellular physiology of renal
organic cation and anion transport. Physiol. Rev. 84, 987-1049.
Zantke, J., Wolff, C. and Scholtz, G. (2008). Three-dimensional reconstruction of the
central nervous system of Macrobiotus hufelandi (Eutardigrada: Parachela): implications
for the phylogenetic position of Tardigrada. Z. Morphol. 127, 21-36.
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Paper III

Acta Physiol 2011, 202, 409–420
REVIEW
Survival in extreme environments – on the current
knowledge of adaptations in tardigrades
N. Møbjerg, 1 K. A. Halberg, 1 A. Jørgensen, 2 D. Persson, 1,2 M. Bjørn, 3 H. Ramløv 3 and
R. M. Kristensen 2
1 Department of Biology, University of Copenhagen, Copenhagen, Denmark
2 Natural History Museum of Denmark, University of Copenhagen, Copenhagen, Denmark
3 Department of Science, Systems and Models, University of Roskilde, Roskilde, Denmark
Received 17 October 2010,
revision requested 13 November
2010,
revision received 6 January 2011,
accepted 10 January 2011
Correspondence: N. Møbjerg,
Department of Biology, August
Krogh Building, Universitetsparken
13, DK-2100 Copenhagen Ø,
Denmark.
E-mail: nmobjerg@bio.ku.dk
Abstract
Tardigrades are microscopic animals found worldwide in aquatic as well as
terrestrial ecosystems. They belong to the invertebrate superclade Ecdysozoa,
as do the two major invertebrate model organisms: Caenorhabditis elegans
and Drosophila melanogaster. We present a brief description of the tardigrades
and highlight species that are currently used as models for physiological
and molecular investigations. Tardigrades are uniquely adapted to a
range of environmental extremes. Cryptobiosis, currently referred to as a
reversible ametabolic state induced by e.g. desiccation, is common especially
among limno-terrestrial species. It has been shown that the entry and exit of
cryptobiosis may involve synthesis of bioprotectants in the form of selective
carbohydrates and proteins as well as high levels of antioxidant enzymes and
other free radical scavengers. However, at present a general scheme of
mechanisms explaining this phenomenon is lacking. Importantly, recent
research has shown that tardigrades even in their active states may be extremely
tolerant to environmental stress, handling extreme levels of ionizing
radiation, large fluctuation in external salinity and avoiding freezing by
supercooling to below )20 °C, presumably relying on efficient DNA repair
mechanisms and osmoregulation. This review summarizes the current
knowledge on adaptations found among tardigrades, and presents new data
on tardigrade cell numbers and osmoregulation.
Keywords cell numbers, cryptobiosis, evolution, osmoreglation, supercooling,
tardigrade.
Tardigrades, also known as water bears, are microscopic
metazoans (approx. 0.1–1.2 mm). They were
discovered in the 18th Century with the development of
early microscopes and were first described by the
German zoologist Goeze in 1773, who named them
‘kleiner Wasserbär’ or little water bear because of their
strong resemblance to a little bear (see e.g. Ramazzotti
& Maucci 1983, Nelson 2001, Schill 2010). Shortly
after in 1776, the current name, Tardigrada (from Latin
tardigradus, slow-moving), was given by the Italian
natural scientist Spallanzani (see e.g. Rebecchi et al.
2007). Tardigrades are exceptional among metazoans in
their adaptations to the most extreme environments. As
is also known from selected species of arthropods,
nematodes and rotifers, many species have the ability to
enter cryptobiosis; a state of suspended animation
believed to be ametabolic (Keilin 1959, Clegg 2001).
Corti already noted these adaptations in tardigrades in
1774, when he observed that these animals could be
revived after desiccation (Kinchin 1994). In 1962,
Tardigrada was recognized as a phylum by Ramazzotti
in Il Phylum Tardigrada (Ramazzotti 1962). There are
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Adaptation to extreme environments in tardigrades Æ
N. Møbjerg et al. Acta Physiol 2011, 202, 409–420
currently approx. 1000 described species (Guidetti &
Bertolani 2005, Degma et al. 2010), but this is likely far
from the real number of tardigrade species, as especially
the marine arthrotardigrades remain relatively unexplored.
Tardigrada belongs to the invertebrate superclade
Ecdysozoa, however; their precise phylogenetic position
is still debated and it is presently not clear whether the
group is more closely related to arthropods and
onychophorans or to the nematodes and nematomorphs
(Fig. 1) (Aguinaldo et al. 1997, Dunn et al. 2008,
Edgecombe 2010). In any case, as noticed in recent
papers, this group has a central position placed in
between the two major invertebrate model organisms –
the nematode Caenorhabditis elegans Maupas, 1900
and the arthropod Drosophila (Sophophora) melanogaster
Meigen, 1830 (Goldstein & Blaxter 2002, Gabriel
& Goldstein 2007).
Little is known about the physiological mechanisms
underlying adaptations to extreme environmental conditions
in tardigrades. Past centuries of tardigrade
research have mainly focused on species descriptions
and morphological investigations related to phylogenetic
analysis. In recent years, however, research in the
field has taken advantage of new molecular tools and an
increasing number of scientists find the group fasci-
Loricifera
Kinorhyncha
Priapulida
ECDYSOZOA
Nematomorpha
Nematoda*
Tardigrada*
Arthropoda*
Onychophora
PROTOSTOMIA
Brachiopoda
Annelida
LOPHOTROCHOZOA
Mollusca
Rotifera*
Platyhelminthes
DEUTEROSTOMIA
Chordata
Echinodermata
Cnidaria
Figure 1 Evolutionary position of tardigrades in the Animal Kingdom. Phylogeny of the Metazoa (animals) based on
Aguinaldo et al. (1997) and Dunn et al. (2008) showing selected phyla with emphasis on the position of the Tardigrada. The
inferred position of the tardigrades is based on EST sequences from Richtersius coronifer and Hypsibius dujardini. Ecdysozoa
includes all molting animals and is one of the two protostome superclades. The marked phyla have cryptobiotic species.
Porifera
410
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Acta Physiol 2011, 202, 409–420
N. Møbjerg et al. Æ
Adaptation to extreme environments in tardigrades
nating. In particular, the mechanisms underlying the
ability to enter cryptobiosis have attracted considerable
scientific interest. This interest is no doubt associated
with the suspected translational output related to a
detailed understanding of the complex stress physiology
of tardigrades, i.e. in connection with cryopreservation
and dehydration of biological material. Noticeably, the
phenomenon cryptobiosis touches upon our conception
of life and death; one of the largest enigmas being how
metabolism is restarted after years of suspension. This
review puts focus on physiological and molecular
adaptations to extreme conditions found among tardigrades.
We additionally present a brief description of
the phylum and highlight species that are currently used
as models for these investigations.
Phylum Tardigrada
Tardigrade phylogeny and evolution
About 35 extant animal groups have body plans and
genes that are distinct enough to warrant elevation to
phylum status (Nielsen 2001). The tardigrades, comprising
the phylum Tardigrada, are one of these groups
(Fig. 1). Tardigrades are microscopic invertebrates with
a well developed organization including brain and
sensory organs, muscles, a complex feeding apparatus
and alimentary tract, reproductive and osmoregulatory
organs (see e.g. Rebecchi & Bertolani 1994, Dewel &
Dewel 1996, Møbjerg & Dahl 1996, Eibye-Jacobsen
1997, Jørgensen et al. 1999, Greven 2007, Halberg
et al. 2009a). They are found worldwide in aquatic as
well as terrestrial environments, but depend on free
water to be in their active, reproducing state. It has been
suggested that tardigrades, like e.g. nematodes, have
eutely, but detailed studies on the subject are still
lacking. Cell counts based on nuclear staining with
DAPI (4¢,6-diamidino-2-phenylindole) in four active
stage adults of the marine eutardigrade Halobiotus
crispae Kristensen, 1982 revealed a total cell number of
around 1060 cells, when excluding gametes (Fig. 2).
This number could represent a slight underestimate of
the total somatic cell number as body cavity cells (also
known as storage cells) may have escaped two of the
specimens, which were punctured prior to the staining
(Fig. 2). We did not observe cell divisions (mitosis)
during the counts. Mitosis has, however, previously
been reported in post-embryonic eutardigrades (Bertolani
1970a,b, 1982).
There are two main evolutionary lines within the
tardigrades, represented by the classes Eutardigrada and
Heterotardigrada (Fig. 3) (see e.g. Jørgensen & Kristensen
2004). The validity of a third class, Mesotardigrada,
is currently uncertain. Mesotardigrada only
contains a single species, Thermozodium esakii Rahm,
1937 originally found in a hot spring in Japan. The type
specimens of T. esakii no longer exist and the type
locality was apparently destroyed in an earthquake
A
P
br
pb
gI
c.gl.
Leg 1
eo
gII
c.gl.
Leg 2
mg
gIII
c.gl.
Leg 3
go
gIV
c.gl.
Leg 4
Figure 2 Cell numbers in Halobiotus crispae. 3-D reconstruction of cell arrangement in the eutardigrade H. crispae based on
confocal laser scanning microscopy of a DAPI stained specimen. In order to obtain an estimate of somatic cell numbers in this
species, specimens were relaxed in CO 2 -enriched water, fixed in 4% paraformaldehyde and ultrasonicated. Two of the four
specimens were additionally delicately punctured with a fine needle. The specimens were subsequently incubated with DAPI and
thoroughly rinsed before mounting. Image acquisition was performed using a Leica DM RXE 6 TL microscope equipped with a
Leica TCS SP2 AOBS confocal laser scanning unit. Cell counts and image processing were performed using the software program
Imaris (Bitplane, Zurich, Switzerland). The total number of somatic cells in H. crispae was estimated at approx. 1060, based on
stainings of four male specimens (mean SD: 1058 53). This number could represent a slight underestimate of the total somatic
cell number as so-called body cavity cells may have escaped the two specimens that were punctured prior to staining (cell counts for
the punctured specimens: 998 and 1088; counts for the non-punctured specimens: 1036 and 1112). The largest number of cells is
clearly present in the anterior part of the animal containing the brain and buccopharyngeal apparatus. A, anterior; P, posterior; br,
brain; c.gl., claw gland; eo, esophagus; gI–IV, ventral ganglia I–IV; mg, midgut; go, gonad; pb, pharyngeal bulb. Scale bar: 50 lm.
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Adaptation to extreme environments in tardigrades Æ
N. Møbjerg et al. Acta Physiol 2011, 202, 409–420
Echiniscoides
Echiniscoidae
Echiniscus
Echiniscidae
Milnesium*
Milnesiidae
Halobiotus
Isohypsibioidea
Richtersius
Paramacrobiotus
Macrobiotoidea
Hypsibius*
Ramazzottius*
Hypsibioidea
Arthrotardigrada
marine
non-cryptobionts
Echiniscoidea
limno-terrestrial
Intertidal
Cryptobionts
Apochela
limno-terrestrial
Cryptobionts
Parachela
limno-terrestrial
cryptobionts
non-cryptobionts
Heterotardigrada
Mesotardigrada ()
Eutardigrada
Tardigrada
*Genome projects
Figure 3 Tardigrade phylogeny. Phylogeny
of tardigrades showing major clades
and position of model/discussed species.
The phylogeny is based on Sands et al.
(2008).
(Nelson 2002). However, a thorough re-sampling for
this species has to our knowledge not been performed.
Tardigrades most likely evolved within the marine
environment, and marine species are especially numerous
within the heterotardigrade order Arthrotardigrada
(Renaud-Mornant 1982, Maas & Waloszek 2001,
Jørgensen et al. 2010). Arthrotardigrades are present
in all oceans from intertidal zones to abyssal depths,
inhabiting different sediment types. In addition, marine
species are found within the other main heterotardigrade
order, Echiniscoidea, represented by the intertidal
Echiniscoides sigismundi (M. Schultze, 1865). This
species may very well be the toughest creature on
Earth, having to endure periods of desiccation and low
oxygen tension as well as large perturbations in salinity
and freezing (Kristensen & Hallas 1980). Nevertheless,
the exact range of this tardigrade’s tolerances remains
to be investigated. It may be hypothesized that an
Echiniscoides-like tardigrade invaded the freshwater/
terrestrial environment and gave rise to the almost
exclusively limno-terrestrial eutardigrades (Kinchin
1994). This is however currently not supported by
molecular data (Sands et al. 2008, Jørgensen et al.
2010; Jørgensen et al. 2011).
The eutardigrades are divided into two orders;
Apochela and Parachela. Two genera within the latter
order, Ramajendas represented by Ramajendas renaudi
(Ramazzotti 1972) in the Southern Hemisphere and
Halobiotus in the Northern Hemisphere, have secondarily
invaded the marine environment (Ramazzotti
1972, Kristensen 1982, Møbjerg et al. 2007). Although
cryptobiosis is common in most eutardigrades, our
recent findings in H. crispae suggest that among these
secondary marine species, adaptations are present that
are quite extraordinary (Halberg et al. 2009b). The
tardigrades stay active while experiencing large fluctuations
in abiotic factors, fluctuations that in other
tardigrades would induce cryptobiosis.
Tardigrade genomes
There is a huge variation in the genome size of
tardigrades ranging from about 75–100 Mb in Hypsibius
and Ramazzottius to 800 Mb in Bertolanius
(Gregory 2010, C-values converted from picograms to
base pairs using the conversion 1 pg = 978 Mb according
to Dolezel et al. (2003), Bertolanius was previously
named Amphibolus). For comparison, the genome sizes
of Caenorhabditis elegans and Drosophila melanogaster
are about 100 Mb and 175 Mb respectively (Gregory
2010). The general diploid chromosome numbers of
the eutardigrades are 10–12 and 14 for the heterotardigrade
Echiniscus (Bertolani 1982). Polyploidy with
up to 24 chromosomes is common in eutardigrades
(Bertolani 1982). Three major sequencing projects are
currently ongoing within Tardigrada. However, all
three projects are investigating eutardigrade species;
no data are presently available for the other main
tardigrade group – the heterotardigrades. The international
collaborative Ecdysozoan Sequencing Project is
assembling the genome of Hypsibius dujardini (Doyère,
1840) as part of an investigation into the ancestral
genome of the Ecdysozoa lineage. Prior to this project,
the Edinburgh based TardiBASE project, generated
more than 5000 EST sequences for H. dujardini
(GenBank 2010). The German based FUNCRYPTA
project was focused on investigating cryptobiosis in
Milnesium tardigradum Doyère, 1840 through studies
of gene and protein expression (Förster et al. 2009,
Mali et al. 2010, Schokraie et al. 2010). The project
had in 2010 generated approx. 7000 quality EST
sequences and aimed at advancing the basic understanding
of protein expression in tardigrades through transcriptomic
and proteomic studies (Mali et al. 2010). The
Japanese based Kumamushi Genome Project is assembling
the genome of Ramazzottius varieornatus Bertolani
and Kinchin, 1993 and has preliminarily predicted
412
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Acta Physiol 2011, 202, 409–420
N. Møbjerg et al. Æ
Adaptation to extreme environments in tardigrades
about 20 000 candidate genes in this species (Horikawa
et al. 2008, Katayama et al. 2009). The great challenge
in the years to come will be to correlate the description of
tardigrade genes with studies on the function of these
genes as nicely illustrated in a recent study on a
purinergic P2X receptor from H. dujardini (Bavan et al.
2009).
Adaptation to extreme environments
Tardigrades are extraordinary in their tolerance to
extremes, including limno-terrestrial habitats that frequently
dry out, habitats that freeze and habitats that
experience large fluctuations in e.g. osmotic pressure
and oxygen tension. Cryptobiosis, referred to as a
reversible ametabolic state induced by unfavorable
environmental conditions, is a common adaptation
especially among limno-terrestrial tardigrade species
(see e.g. Wright 2001). Four cryptobiosis inducing
physical extremes are traditionally recognized: dehydration
(anhydrobiosis), extremely low temperatures
(cryobiosis), lack of oxygen (anoxybiosis) and high salt
concentration (osmobiosis) (Keilin 1959), with desiccation
induced anhydrobiosis and freezing induced cryobiosis
being the most extensively studied states.
Anhydrobiosis and cryobiosis are not equivalent phenomena
and likely involve different mechanisms for
protection of cells and tissues (Crowe et al. 1990,
1992). Little is known of cryobiosis in tardigrades
(Westh et al. 1991, Ramløv & Westh 1992, Westh &
Kristensen 1992, Halberg et al. 2009b, Hengherr et al.
2009, 2010), whereas a great deal of attention has been
paid to anhydrobiosis (Wright et al. 1992, Wright
2001, Rebecchi et al. 2007, Schill 2010). The anhydrobiotic
state is characterized by the formation of a socalled
tun with withdrawn legs and a longitudinally
contracted body (see e.g. Bertolani et al. 2004). Tun
formation is also seen in bdelloid rotifers, whereas
desiccation tolerant nematodes coil into a tight spiral.
Obviously, the ability to pack internal organs during
tun formation is an important adaptation to desiccation.
Importantly, tun formation is an active, regulated
event and not merely an effect of water removal (Crowe
1972). Along this line, our unpublished data show that
when we expose the active state of the cryptobiotic
tardigrade Richtersius coronifer (Richters, 1903) to
water containing high levels of chemical substances, the
tardigrades will respond by contracting their bodies
initiating tun formation thus undergoing chemobiosis–a
cryptobiotic response to environmental toxins. The
formation of the tun is a critical and necessary event
for tardigrades entering anhydrobiosis. Much more
work needs to be done in order to understand the
processes undertaken during transformation to this
ametabolic state, from the molecular to whole organism
level, anatomically as well as physiologically. In the tun
state tardigrades may not only stand long periods of
desiccation and exposure to toxic chemicals but also
very low subzero temperatures, vacuum, high pressure,
radiation, extreme pH, anoxia and to some extent high
temperature (see e.g. Wright 2001, Rebecchi et al.
2007). It has been suggested and to some degree shown
that adaptations to these extremes may involve synthesis
of bioprotectants in the form of selective carbohydrates
and proteins, high levels of antioxidant enzymes
and other free radical scavengers, biological membranes
containing specific phospholipids as well as powerful
DNA repair mechanisms (Westh & Ramløv 1991, Schill
et al. 2004, Jönsson et al. 2005, Rizzo et al. 2010).
In addition to cryptobiotic tardigrade species, we also
find species that form cysts and enter diapause (see e.g.
Møbjerg et al. 2007, Guidetti et al. 2008). Importantly,
it has been shown that tardigrades even in their active
states may be extremely tolerant to environmental stress
(May et al. 1964, Jönsson et al. 2005, Horikawa et al.
2006, Halberg et al. 2009b). Virtually nothing is known
about the normal physiology of tardigrades, and
answers to how they tolerate these extremes may
actually be found here. We have recently shown that
the littoral eutardigrade Halobiotus crispae handles
large fluctuation in external salinity and avoids freezing
by supercooling to around )20 °C in its active stage
(Halberg et al. 2009b). This species is characterized by
seasonal cyclic changes in morphology and physiology
known as cyclomorphosis, one of the cyclomorphic
stages being freeze tolerant (Kristensen 1982, Møbjerg
et al. 2007, Halberg et al. 2009b). The morphological
changes occurring during cyclomorphosis in H. crispae
in some respects resemble the formation of dauer larvae
in C. elegans (see e.g. Cassada & Russell 1975, Burnell
et al. 2005).
Cryptobiosis in tardigrades
Many experiments on tardigrade cryptobiosis have been
performed on the eutardigrade Richtersius coronifer.
R. coronifer, also known as the giant yellow water bear,
has a body length of up to 1 mm (Fig. 4a–c). Both males
and females are present in some populations, but, as
commonly found among eutardigrades, several populations
reproduce by parthenogenesis. R. coronifer lives in
moss in alpine and arctic environments and is furthermore
numerous in moss on carbonated bedrock in dry
Swedish lowland areas known as Alvar (see e.g. Westh
& Kristensen 1992, Jönsson et al. 2001). R. coronifer is
a true cryptobiont, tolerating extreme desiccation as
imposed by e.g. space vacuum conditions (Jönsson et al.
2008, Persson et al. 2011) and it additionally survives
exposures to very low temperatures as encountered by
transfers into liquid nitrogen (approx. )196 °C) in the
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Adaptation to extreme environments in tardigrades Æ
N. Møbjerg et al. Acta Physiol 2011, 202, 409–420
(a) (b) (c)
st
pb
eo
A
P
A
P
A
P
mg
st
pb mg
*
*
*
(d) (e) (f)
st
st
st
pb
eo
pb
eo
pb
mg
mg
mg
go
* * * *
*
* * *
*
Figure 4 Cryptobiotic and non-cryptobiotic survival in extreme environments. (a–c): Microscopy of Richtersius coronifer from
Stora Alvar on the Swedish island Öland in the Baltic Sea. Richtersius coronifer has been a model tardigrade for investigations on
cryptobiotic survival. (a) Active state; (b) light microscopy and (c) scanning electron microscopy of the cryptobiotic tun state
induced by desiccation. In this state R. coronifer e.g. tolerates complete desiccation as experienced by vacuum condition and
freezing in liquid nitrogen. (d–f): Microscopy of Halobiotus crispae from the Danish population at Vellerup Vig, Denmark. This
tardigrade is one of a few species of eutardigrades that have secondarily invaded the marine environment. Halobiotus crispae in the
active stage (d) uniquely adapted to cope with profound changes in ambient salinity occurring in tidal and subtidal habitats. Upon
transfers to dilute salt water solutions, active stage H. crispae swell (e) and subsequently regulate their body volume to near control
conditions. Halobiotus crispae is characterized by appearing in different cyclomorphic stages. The pseudosimplex 1 stage (f) is freeze
tolerant. eo, esophagus; mg, midgut; pb, pharyngeal bulb; st, stylet. Asterisks mark the position of three Malpighian tubules
presumably involved in osmoregulation. Scale bars: 50 lm.
tun as well as active hydrated state, however,with the
tuns tolerating considerable longer time of exposure
(Ramløv & Westh 1992, Persson et al. 2011). The
anhydrobiotic state survives temperatures of up to
approx. 70 °C for 1 h, but survival rapidly decreases
when the temperature exceeds 70 °C, and no specimens
survives exposure to 100 °C (Ramløv & Westh 2001).
The increase in life span offered by anhydrobiosis in this
species seems to be restricted to approx. 5 years (Westh
& Kristensen 1992). This is less than the cryptobiotic
life expansion observed for the heterotardigrade Echiniscus
testudo (Doyère, 1840), for which we have
revived specimens from moss cushions dried for approx.
20 years (Jørgensen et al. 2007). What then sets the
limits for cryptobiotic survival Probably accumulated
damage to DNA and other molecules as well as to
tissues and organs obtained during the ametabolic state.
This includes damage obtained through oxidative processes
as well as predation, bacterial and fungal
infections. A recent investigation in Paramacrobiotus
richtersi (Murray, 1911) has shown that regulation of
antioxidant metabolism likely plays an important role
in defense against the potential oxidative damage
associated with dehydration (Rizzo et al. 2010). It has
long been known that there is a positive correlation
between the time spent in the anhydrobiotic state and
the time required for recovering active life after rehydration
(Crowe & Higgins 1967). Recent investigations
indicate that the longer the time spent in anhydrobiosis,
the more damage is inflicted to DNA (Neumann et al.
2009, Rebecchi et al. 2009b), which would explain the
prolonged recovery time, indicating that considerable
repair of DNA (and other molecules) occurs in the postanhydrobiotic
state.
Trehalose accumulation. Current knowledge suggests
that specific carbohydrates and proteins are important
in protecting the cells from damage encountered during
414
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Acta Physiologica Ó 2011 Scandinavian Physiological Society, doi: 10.1111/j.1748-1716.2011.02252.x

Acta Physiol 2011, 202, 409–420
N. Møbjerg et al. Æ
Adaptation to extreme environments in tardigrades
entry and exit of cryptobiosis. Bioprotectants may
interact directly with macromolecular structures such
as membranes, DNA and proteins, or they may act as
osmolytes during osmotic or dehydration stress. As has
been shown for other cryptobiotic invertebrates, noticeably
in the cysts of the brine shrimp Artemia franciscana
Kellogg, 1906, some, but not all, cryptobiotic
tardigrades accumulate the dissacharide trehalose during
desiccation (Clegg 1965, Westh & Ramløv 1991,
Hengherr et al. 2008, Jönsson & Persson 2010). Trehalose
has been proposed to act as a molecular stabilizer
replacing water and to further stabilize cellular structure
through the formation of amorphous glasses, a
process known as vitrification (Clegg 2001). The highest
trehalose concentrations measured in anhydrobiotic
tardigrades ranges from 2.3% d.w. in R. coronifer
and Macrobiotus krynauwi Dastych and Harris, 1995
to 2.9% d.w. in Macrobiotus islandicus Richters, 1904
(Westh & Ramløv 1991, Jönsson & Persson 2010).
These trehalose values are, however, relatively low
when compared with the above mentioned crustacean,
which accumulates the disaccharide in concentrations of
around 15% d.w. Moreover, trehalose levels are barely
measurable and do not increase during dehydration in
the cryptobiotic Milnesium tardigradum (see Hengherr
et al. 2008, Jönsson & Persson 2010). The latter
investigations, analysing trehalose contents in eutardigrades
as well as heterotardigrades, show that
trehalose accumulation does not represent a universal
protective mechanism enabling tardigrades to undergo
cryptobiosis.
Expression of heat-shock proteins. Cryptobiosis has
been suggested to rely on the synthesis of molecular
chaperones such as heat-shock-proteins, which may
assist folding of newly synthesized proteins, control
their final intracellular location, as well as protect them
from stress-associated denaturation and aid in renaturation
(Clegg 2001). During entrance into anhydrobiosis
Ramløv & Westh (2001) described the upregulation of
a protein with a molecular weight of approx. 71 kDa in
R. coronifer and proposed that this protein might
belong to the heat-shock-protein (Hsp70) family. Schill
et al. (2004) subsequently investigated RNA expression
patterns of three Hsp70 isoforms in active and anhydrobiotic
states of M. tardigradum as well as during
entrance into and exit out of anhydrobiosis. Only one
of these isoforms (isoform 2) was significantly induced
by the transition from the active to cryptobiotic state
and interestingly, it showed a considerable increase in
expression during the post-cryptobiotic phase, while
the other isoforms were down regulated during cryptobiosis
as well as in transitional states. Jönsson &
Schill (2007) used an immuno-westernblot method to
quantify the induction of Hsp70 in R. coronifer in
response to desiccation, ionizing radiation and heating.
They reported elevated levels of Hsp70 following both
heat and radiation treatment as well as in tardigrades
rehydrated after a period of desiccation. Noticeably,
the authors found that tardigrades in the desiccated
state had reduced Hsp70 levels as compared to the nontreated
control group and accordingly suggested that
Hsp70 may be involved in the repair processes after
desiccation rather than acting as a biochemical stabilizer
in the dry state. Based on the M. tardigradum EST
library, several additional heat-shock proteins have
been identified, including two a-crystalline heat-shock
proteins, and the relative abundance of the transcripts
coding for these stress proteins have been investigated
during phases of dehydration and rehydration (Reuner
et al. 2010). The results obtained suggested a limited
role for heat-shock proteins in the desiccation tolerance
of M. tardigradum. The authors found a variable
pattern of expression with most of the candidate genes
being down regulated, and only one of the genes (Mthsp90),
being significantly upregulated in the dehydrated
state (Reuner et al. 2010). Comparable studies
in Paramacrobiotus richtersi did not show evidence for
an increased expression of either Hsp70 or Hsp90
between hydrated and dehydrated animals (Rizzo et al.
2010). Additionally, specimens of P. richtersi sent into
space (Foton-M3 mission) revealed no significant
change as compared to ground controls in the expression
of these heat-shock proteins (Rebecchi et al.
2009a). These contrasting results indicate that Hsp
expression is species specific. A general role in the
cryptobiotic survival of tardigrades can at present not
be attributed to these stress proteins. Nevertheless, a
synergistic effect between HSPs and other bioprotectants
such as trehalose or LEA (late-embryogenesis
abundant) proteins might exist (see e.g. Goyal et al.
2005). Notably, LEA proteins have been described in
many other desiccation tolerant organisms from plants
to arthropods and might analogously be functionally
important in tardigrades (see e.g. Schill 2010, Warner
et al. 2010). Indeed, a putative LEA protein has
recently been detected in M. tardigradum (Schokraie
et al. 2010). Consequently, future studies on these and
other bioprotectants might offer additional clues in our
search to understand the phenomenon cryptobiosis.
Obviously, cryptobiosis is far from understood and
much more research is needed in order to understand
the mechanisms underlying this state. Interestingly, a
preliminary study from the Kumamushi Genome
Project on R. varieornatus suggests that comparative
metabolome profiling of active and anhydrobiotic
states may provide a novel insight into anhydrobiosis
(Arakawa et al. 2009). Additionally, clues to cryptobiotic
survival may be found in the extreme tolerance
seen in active state tardigrades.
Ó 2011 The Authors
Acta Physiologica Ó 2011 Scandinavian Physiological Society, doi: 10.1111/j.1748-1716.2011.02252.x 415

Adaptation to extreme environments in tardigrades Æ
N. Møbjerg et al. Acta Physiol 2011, 202, 409–420
Non-cryptobiotic survival in tardigrades
While tardigrades are well known for their abilities to
cope with extreme environmental conditions by entering
cryptobiosis, little focus has been on their ability to
sustain metabolism and remain active during fluctuating
external circumstances. In this context it is interesting to
note that tardigrades are genuine aquatic animals – they
are dependent on free water to be in their active feeding
and reproducing states. Truly terrestrial conditions are
only survived following entry into the cryptobiotic
state. Furthermore, limno-terrestrial tardigrades that
have the ability to enter the tun state may do so in
response to environmental challenges (e.g. exposure to
chemical substances) that in marine species would not
force the animals into cryptobiosis. In the following we
discuss what is currently known about the physiology
and stress tolerance of tardigrades in their active states.
Radiation tolerance. Investigations on radiation tolerance
in Richtersius coronifer have revealed that exposure
to c-radiation at doses up to 1 kGy does not affect
survival of desiccated nor hydrated animals, with
hydrated animals tolerating doses of up to 5 kGy (Jönsson
et al. 2005). Horikawa and co-workers revealed that
both hydrated and desiccated Milnesium tardigradum
survive doses of c-radiation of more than 5 kGy, and up to
8 kGy of heavy ion radiation (Horikawa et al. 2006).
These and previous studies indicate that hydrated animals
are just as good or even better at tolerating radiation. This
indicates that radiation tolerance is not due to biochemical
protectants associated with the cryptobiotic state, but
suggests that tardigrades rely on efficient and yet unidentified
mechanisms of DNA repair (Jönsson et al. 2005,
Horikawa et al. 2006). Several recent studies involved
with the BIOPAN 6/Foton-M3 mission in 2007, funded
by the European Space Agency, have investigated the
impact on survival of tardigrades exposed to space
conditions (Jönsson et al. 2008, Rebecchi et al. 2009a,
Persson et al. 2011). During this mission, cryptobiotic
tardigrades (as well as nematodes and rotifers) were sent
into low earth orbit and exposed to space vacuum and
cosmic radiation. The three studies revealed discrepancies
in survival rates likely reflecting differences between
tardigrade species and experimental setups, however;
they unanimously conclude that tardigrades can survive
the rigors of space, and that M. tardigradum is likely the
most resistant species with embryos as well as adults
tolerating space conditions (see discussion in Persson
et al. 2011). Detailed knowledge of the life history of this
predatory tardigrade is available as a result of a comprehensive
study by Suzuki (2003).
Osmoregulation. Experiments on the marine eutardigrade
Halobiotus crispae have revealed an extraordinary
tolerance to perturbations in body volume and osmotic
pressure (Halberg et al. 2009b). The tidal habitat in
which this tardigrade lives is characterized by large
fluctuations in abiotic factors; most noticeably, alterations
are seen in salinity and temperature (Møbjerg et al.
2007). One way of coping with these extremes would be
to enter cryptobiosis. However, H. crispae is not a
cryptobiont. In the active stage the tardigrade handles
extremes by expending energy on active regulatory
mechanisms. In addition, the so-called pseudosimplex 1
(P1) stage of this animal is freeze tolerant (Halberg et al.
2009b); it is not, however, a cryobiont as the P1 stage does
not tolerate gradual freezing to )80 °C. Cryobiosis is
defined by the apparent absence of a lower lethal
temperature (Wright 2001). The P1 stage is distinctly
characterized by a double cuticle and closed mouth and
cloaca (Kristensen 1982, Møbjerg et al. 2007).
Halobiotus crispae kept at a salinity of 20 ppt have an
extensive ability to supercool (avoid freezing) down to
around )20 °C, enabling active stage animals to withstand
subzero temperatures without freezing (Halberg
et al. 2009b). Similar supercooling points have been
reported from limno-terrestrial eutardigrades (Macrobiotus,
Paramacrobiotus and Milnesium), whereas heterotardigades
(Echiniscus) had slightly higher points
(Hengherr et al. 2009). Much higher supercooling
points of )6.7 °C and )7.4 °C were originally observed
in respectively Richtersius coronifer and Bertolanius
nebulosus (Westh & Kristensen 1992). These high
supercooling points are likely the result of the presence
of ice-nucleating agents initiating the freezing process
(Westh et al. 1991). Figure 5 shows thermograms of H.
crispae kept in seawater with a salinity of 20 ppt
(Fig. 5a) and distilled water (Fig. 5b) prior to differential
scanning calorimetry (DSC). Water content in the
animals kept at 20 ppt and in distilled water was
respectively 73% and 81%. The latter group of tardigrades
had a higher supercooling point as compared to
the group kept in 20 ppt saltwater, illustrating the
expected correlation between water content and crystallization
temperature in species without ice-nucleating
agents (see e.g. Hengherr et al. 2009). Thus, the extent
of supercooling and thereby the ability to avoid freezing,
is coupled to osmoregulation.
Halobiotus crispae has a large capacity to tolerate
perturbations in ambient salinity making it an ideal
model for the study of osmoregulation and volume
regulation in tardigrades. Experiments on this species
revealed tolerance to a wide range of salinities, with
specimens of the Greenlandic population remaining
active in solutions ranging from distilled water to
saltwater with osmolalities up to 2000 mOsm kg )1
(Halberg et al. 2009b). During experiments with transfers
to strong hypotonic solutions, active stage H.
crispae swell with up to 60% before regulating back to
416
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Acta Physiol 2011, 202, 409–420
N. Møbjerg et al. Æ
Adaptation to extreme environments in tardigrades
(a) 15
(a) 1400
Differential heat flow (mW)
Differential heat flow (mW)
7.5
0
(b) 15
7.5
0
Exotherm Exotherm
Cooling
Heating
near control values (Fig. 4e). Similarly, specimens
transferred into hypertonic solutions shrink and thereafter
respond by regulating body volume. In a series of
experiments with transfers into saltwater solutions with
a salinity between 2 ppt (62 mOsm kg )1 ) and 40 ppt
(1245 mOsm kg )1 ) the active stage tardigrades hyperregulated
at all times (Fig. 6a) (Halberg et al. 2009b).
Hyperregulation is likely a general feature of at least the
eutardigrades, as our data on R. coronifer reveal that
this species also keeps its body fluids hyperosmotic as
compared to the surroundings (Fig. 6b). Our data
indicate that this limno-terrestrial cryptobiont is less
tolerant of high salinity solutions than the littoral H.
crispae. When exposed to water with increasing salt
Endotherm
20 ppt.
–40 –35 –30 –25 –20 –15 –10 –5 0 5
Cooling
Heating
Endotherm
0 ppt.
–40 –35 –30 –25 –20 –15 –10 –5 0 5
Temperature (°C)
Figure 5 Supercooling in Halobiotus crispae. (a) Supercooling
in H. crispae kept in seawater with a salinity of 20 ppt.
Thermogram modified from Halberg et al. (2009b). (b) Halobiotus
crispae kept in distilled water. Cooling and subsequent
reheating of a sample containing 46 specimens of H. crispae
from Vellerup Vig, Denmark kept in distilled water for 45 min
prior to the transfer into pans for differential scanning calorimetry
(DSC). Freezing ()12.2 °C) and melting ()1.2 °C)
temperatures were estimated as onsets of peaks of respectively
exothermic and endothermic events using DSC7 software
(cooling rate 2 °C min )1 ) (DSC7; Perkin Elmer Inc., Wellesley,
MA, USA).
Hemolymph osmolality (mOsm kg –1 ) Hemolymph osmolality (mOsm kg –1 )
1200
1000
800
600
400
200
(b) 1400
1200
1000
H. crispae
Osmotic performance
Isoosmotic line
0
0 200 400 600 800 1000 1200 1400
800
600
400
200
R. coronifer
Osmotic performance
Isoosmotic line
0
0 200 400 600 800 1000 1200 1400
External Osmolality (mOsm kg –1 )
Figure 6 Osmoregulation in eutardigrades. (a) Measured hemolymph
osmolality of active stage Halobiotus crispae from
Vellerup Vig, Denmark as a function of external osmolality.
Figure modified from Halberg et al. (2009b). (b) Hemolymph
osmolality of Richtersius coronifer from Öland, Sweden as a
function of external osmolality. External solutions with osmotic
concentrations of 100, 200, 300 and 500 mOsm kg )1 were
prepared from artificial seawater salt (Tropic Marin, Dr Biener
GmbH, Germany). Each point on the graph represents
mean SD of hemolymph osmolality measurements (nanolitre
osmometry; Clifton Technical Physics, Hartford, NY, USA)
made in tardigrades kept in demineralized water and the different
salt water solution for 30 min. Five animals were used for
hemolymph determination at each of the experimental solutions.
Arrow indicates the upper lethal line for R. coronifer.
concentrations, R. coronifer will become inactive and
eventually die, exhibiting an upper lethal limit of
around 500 mOsm kg )1 (Fig. 6b). In solutions ranging
from demineralized water to salt water with an osmolality
of 500 mOsm kg )1 , the tardigrade, however,
maintains a consistent osmotic gradient of around
170 mOsm kg )1 above that of the external environment
(Fig. 6b). In comparison, H. crispae exposed to the
same salinity range maintains an osmotic gradient of
around 300 mOsm kg )1 above that of the surroundings
(Fig. 6a). These data on two different species of
tardigrades indicate that eutardigrades have a relatively
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Adaptation to extreme environments in tardigrades Æ
N. Møbjerg et al. Acta Physiol 2011, 202, 409–420
high water turnover, and that they excrete a hypoosmotic
fluid – the likely organs involved in this
excretion being Malpighian tubules and the gut system
(Fig. 4) (Møbjerg & Dahl 1996, Halberg et al. 2009b).
At present the composition of tardigrade extracellular
fluids (as well as intracellular fluids) are unknown.
Obviously, information on the composition of the
hemolymph is much needed for our understanding of
tardigrade physiology and future discussions on how
tardigrades regulate their body fluids and especially
which osmolytes they are regulating. It is likely that the
osmotic pressure obtained through hyperregulation is
partly built by organic solutes, which in turn may act as
bioprotectants or may be important for e.g. the above
mentioned ability to supercool.
In summary, recent research has shown that tardigrades
in their active state may tolerate large fluctuations
in abiotic factors and cope with environmental
alterations by staying active and maintaining high
metabolic rates. It would seem that these extraordinary
organisms have two ways of handling environmental
extremes; one way is entering a dormant state, i.e.
cryptobiosis or diapause/encystment and the other,
likely requiring a high metabolic rate, is relaying on
e.g. osmoregulation and DNA repair, while staying
active. The mechanisms enabling tardigrades to withstand
environmental extremes in their active state may
provide clues to the much debated mechanism underlying
the phenomenon of cryptobiosis. Intriguingly,
adaptations to some of the harshest environments on
Earth have given tardigrades the ability to survive
conditions that by far exceed the extremes presented by
the environment (e.g. extreme levels of ionizing radiation
and extremely low temperatures), presenting an
unresolved puzzle for contemporary biology.
Conflict of interest
There are no conflicts of interest.
We would like to thank Peter Westh for help in generating the
DSC data presented in Figure 5. Special thanks are due to the
organizers of the August Krogh Symposium 2010: Erik Hviid
Larsen, Ylva Hellsten and Jørgen Wojtaszewski. Funding came
from the 2008 Faculty of Science, University of Copenhagen
Freja-Programme and from the Carlsberg Foundation.
References
Aguinaldo, A.M.A., Turbeville, J.M., Linford, L.S., Rivera,
M.C., Garey, J.R., Raff, R.A. & Lake, J.A. 1997. Evidence
for a clade of nematodes, arthropods and other moulting
animals. Nature 387, 489–493.
Arakawa, K., Ito, T., Kunieda, T., Horikawa, D., Soga, T. &
Tomita, M. 2009. Comparative metabolome profiling of
active and anhydrobiotic states of tardigrade Ramazzottius
varieornatus. 11th International Symposium on Tardigrada.
Tübingen, Germany. 3–6 August 2009, Conference Guide, p.
37 (Abstract).
Bavan, S., Straub, V.A., Blaxter, M.L. & Ennion, S.J. 2009. A
P2X receptor from the tardigrade species Hypsibius dujardini
with fast kinetics and sensitivity to zinc and copper.
BMC Evol Biol 9, 17.
Bertolani, R. 1970a. Mitosi somatiche e costanza cellulare
numerica nei Tardigradi. Atti Accad Naz Lincei Rend Ser 8a,
739–742.
Bertolani, R. 1970b. Variabilità numerica cellulare in alcuni
tessuti di Tardigradi. Atti Accad Naz Lincei Rend Ser 8a,
442–445.
Bertolani, R. 1982. Cytology and reproductive mechanisms in
tardigrades. In: D.R. Nelson (ed.) Proceedings of the Third
International Symposium on the Tardigrada, pp. 93–114.
East Tennessee State University Press, Johnson City, TN.
Bertolani, R., Guidetti, R., Jönsson, K.I., Altiero, T., Boschini,
D. & Rebecchi, L. 2004. Experiences with dormancy in
tardigrades. J Limnol 63, 16–25.
Burnell, A.M., Houthoofd, K., O’Hanlon, K. & Vanfleteren, J.R.
2005. Alternate metabolism during the dauer stage of the
nematodeCaenorhabditiselegans.ExpGerontol40,850–856.
Cassada, R.C. & Russell, R.L. 1975. The dauerlarva, a postembryonic
developmental variant of the nematode Caenorhabditis
elegans. Dev Biol 46, 326–342.
Clegg, J.S. 1965. The origin of trehalose and its significance
during formation of encysted dormant embryos of Artemia
salina. Comp Biochem Physiol 14, 135–143.
Clegg, J.S. 2001. Cryptobiosis – a peculiar state of biological
organization. Comp Biochem Physiol B 128, 613–624.
Crowe, J.H. 1972. Evaporative water loss by tardigrades under
controlled relative humidity. Biol Bull 142, 407–416.
Crowe, J.H. & Higgins, R.P. 1967. The revival of Macrobiotis
aereolatus Murray (Tardigrada) from the cryptobiotic state.
Trans Am Microsc Soc 86, 286–294.
Crowe, J.H., Carpenter, J.F., Crowe, L.M. & Anchordoguy,
T.J. 1990. Are Freezing and dehydration similar stress vectors
– a comparison of modes of interaction of stabilizing
solutes with biomolecules. Cryobiology 27, 219–231.
Crowe, J.H., Hoekstra, F.A. & Crowe, L.M. 1992. Anhydrobiosis.
Annu Rev Physiol 54, 579–599.
Degma, P., Bertolani, R. & Guidetti, R. 2010. Actual checklist
of Tardigrada species (Ver. 13: 01-06-2010). http://
www.tardigrada.modena.unimo.it.
Dewel, R.A. & Dewel, W.C. 1996. The brain of Echiniscus viridissimus
Peterfi, 1956 (Heterotardigrada): a key to understanding
the phylogenetic position of tardigrades and the
evolution of the arthropod head. Zool J Linn Soc 116, 35–49.
Dolezel, J., Bartoš, J., Voglmayr, H. & Greilhuber, J. 2003.
Nuclear DNA content and genome size of trout and human.
Cytometry 51A, 127–128.
Dunn, C.W., Hejnol, A., Matus, D.Q., Pang, K., Browne, W.E.,
Smith, S.A., Seaver, E., Rouse, G.W., Obst, M., Edgecombe,
G.D. et al. 2008. Broad phylogenomic sampling improves
resolution of the animal tree of life. Nature 452, 745–750.
Edgecombe, G.D. 2010. Arthropod phylogeny: an overview
from the perspective of morphology, molecular data and the
fossil record. Arthropod Struct Dev 39, 74–87.
418
Ó 2011 The Authors
Acta Physiologica Ó 2011 Scandinavian Physiological Society, doi: 10.1111/j.1748-1716.2011.02252.x

Acta Physiol 2011, 202, 409–420
N. Møbjerg et al. Æ
Adaptation to extreme environments in tardigrades
Eibye-Jacobsen, J. 1997. Development, ultrastructure and
function of the pharynx of Halobiotus crispae Kristensen,
1982 (Eutardigrada). Acta Zool (Stock) 78, 329–347.
Förster, F., Liang, C., Shkumatov, A., Beisser, D., Engelmann,
J.C., Schnölzer, M., Frohme, M., Müller, T., Schill, R.O. &
Dandekar, T. 2009. Tardigrade workbench: comparing
stress-related proteins, sequence-similar and functional protein
clusters as well as RNA elements in tardigrades. BMC
Genomics 10, 469.
Gabriel, W.N. & Goldstein, B. 2007. Segmental expression of
Pax3/7 and Enrailed homologs in tardigrade development.
Dev Genes Evol 217, 421–433.
Goldstein, B. & Blaxter, M. 2002. Tardigrades. Curr Biol 12,
R475.
Goyal, K., Walton, L.J. & Tunnacliffe, A. 2005. LEA proteins
prevent protein aggregation due to water stress. Biochem J
388, 151–157.
Gregory, T.R. 2010. Animal Genome Size Database. http://
www.genomesize.com.
Greven, H. 2007. Comments on the eyes of tardigrades.
Arthropod Struct Dev 36, 401–407.
Guidetti, R. & Bertolani, R. 2005. Tardigrade taxonomy: an
updated check list of the taxa and a list of characters for their
identification. Zootaxa 845, 1–46.
Guidetti, R., Boschini, D., Altiero, T., Bertolani, R. & Rebecchi,
L. 2008. Diapause in tardigrades: a study of factors involved
in encystment. J Exp Biol 211, 2296–2302.
Halberg, K.A., Persson, D., Møbjerg, N., Wanninger, A. &
Kristensen, R.M. 2009a. Myoanatomy of the marine tardigrade
Halobiotus crispae (Eutardigrada: Hypsibiidae).
J Morph 270, 996–1013.
Halberg, K.A., Persson, D., Ramløv, H., Westh, P., Kristensen,
R.M. & Møbjerg, N. 2009b. Cyclomorphosis in Tardigrada:
adaptation to environmental constraints. J Exp Biol 212,
2803–2811.
Hengherr, S., Heyer, A.G., Köhler, H.R. & Schill, R.O. 2008.
Trehalose and anhydrobiosis in tardigrades – evidence for
divergence in response to dehydration. FEBS 275, 281–288.
Hengherr, S., Worland, M.R., Reuner, A., Brümmer, F. & Schill,
R.O. 2009. Freeze tolerance, supercooling pionts and ice formation:
comparative studies on the subzero temperature survival
of limno-terrestrial tardigrades. J Exp Biol 212, 802–807.
Hengherr, S., Reuner, A., Brümmer, F. & Schill, R.O. 2010. Ice
crystallization and freeze tolerance in embryonic stages of
the tardigrade Milnesium tardigradum. Comp Biochem
Physiol A Mol Integr Physiol 156, 151–155.
Horikawa, D.D., Sakashita, T., Katagiri, C., Watanabe, M.,
Kikawada, T., Nakahara, Y., Hamada, N., Wada, S.,
Funayama, T., Higashi, S., Kobayashi, Y., Okuda, T. &
Kuwabara, M. 2006. Radiation tolerance in the tardigrade
Milnesium tardigradum. Int J Radiat Biol 82, 843–848.
Horikawa, D.D., Kuenida, T., Abe, W., Watanabe, M., Nakahara,
Y., Yukuhiro, F., Sakashita, T., Hamada, N., Wada, S.,
Funayama, T., Katagiri, C., Kobayashi, Y., Higashi, S. &
Okuda, T. 2008. Establishment of a rearing system of the extremotolerant
tardigrade Ramazzottius varieornatus: a new
model animal for astrobiology. Astrobiology 8, 549–556.
Jönsson, K.I. & Persson, O. 2010. Trehalose in three species of
desiccation tolerant tardigrades. Open Zool J 3, 1–5.
Jönsson, K.I. & Schill, R. 2007. Induction of Hsp70 by desiccation,
ionising radiation and heat-shock in the eutardigrade
Richtersius coronifer. Comp Biochem Physiol B 146,
456–460.
Jönsson, K.I., Borsari, S. & Rebecchi, L. 2001. Anhydrobiotic
survival in populations of the tardigrades Richtersius coronifer
and Ramazzottius oberhaeuseri from Italy and Sweden.
Zool Anz 240, 419–423.
Jönsson, K.I., Harms-Ringdahl, M. & Torudd, J. 2005. Radiation
tolerance in the eutardigrade Richtersius coronifer. Int
J Radiat Biol 81, 649–656.
Jönsson, I.K., Rabbow, E., Schill, R.O., Harms-Ringdahl, M.
& Rettberg, P. 2008. Tardigrades survive exposure to space
in low Earth orbit. Curr Biol 18, R729–R731.
Jørgensen, A., Faurby, S., Hansen, J.G., Møbjerg, N. & Kristensen,
R.M. 2010. Molecular phylogeny of Arthrotardigrada
(Tardigrada). Mol Phylogenet Evol 54, 1006–1015.
Jørgensen, A. & Kristensen, R.M. 2004. Molecular phylogeny
of Tardigrada – the monophyly of Heterotardigrada. Mol
Phylogenet Evol 32, 666–670.
Jørgensen, A., Møbjerg, N. & Kristensen, R.M. 1999. Ultrastructural
studies on spermiogenesis and postcopulatory
modifications of spermatozoa of Actinarctus doryphorus
Schulz, 1935 (Arthrotardigrada: Halechiniscidae). Zool Anz
238, 235–257.
Jørgensen, A., Møbjerg, N. & Kristensen, R.M. 2007. A
molecular study of the tardigrade Echiniscus testudo (Echiniscidae)
reveals low DNA sequence diversity over a large
geographical area. J Limnol 66(Suppl. 1), 77–83.
Jørgensen, A., Møbjerg, N. & Kristensen, R.M. 2011. Phylogeny
and evolution of the Echiniscidae (Echiniscoidea,
Tardigrada) – an investigation of the congruence between
molecules and morphology. J Zool Syst Evol Res: doi:
10.1111/j.1439-0469.2010.00592.x
Katayama, T., Arakawa, K., Hasebe, Y., Kido, N., Kunieda,
T., Toyoda, A., Shin-I, T., Horikawa, D.D., Kuwahara, H.,
Ohishi, K., Motoyama, A., Aizu, T., Kanehisa, M., Kohara,
Y. & Fujiyama, A. 2009. Draft genome sequence assembly
and preliminary annotations of Ramazzottius varieornatus
genome. 11th International Symposium on Tardigrada.
Tübingen, Germany. 3–6 August 2009, Conference Guide, p.
36 (Abstract).
Keilin, D. 1959. The problem of anabiosis or latent life: history
and current concept. Proc Roy Soc Lond (Ser B) 150, 149–
191.
Kinchin, I.M. 1994. The Biology of Tardigrades, p. 186.
Portland Press, London.
Kristensen, R.M. 1982. The first record of cyclomorphosis in
Tardigrada based on a new genus and species from Arctic
meiobenthos. J Zool Syst Evol Res 20, 249–270.
Kristensen, R.M. & Hallas, T.E. 1980. The tidal genus Echiniscoides
and its variability with erection of Echiniscoididae
fam.n. (Tardigrada). Zool Scr 9, 113–127.
Maas, A. & Waloszek, D. 2001. Cambrian derivatives of the
early arthropod stem lineage, pentastomids, tardigrades and
lobopodians – an ‘Orsten’ perspective. Zool Anz 240, 451–
459.
Mali, B., Grohme, M.A., Förster, F., Dandekar, T., Schnölzer,
M., Reuter, D., Wełnicz, W., Schill, R.O. & Frohme, M.
Ó 2011 The Authors
Acta Physiologica Ó 2011 Scandinavian Physiological Society, doi: 10.1111/j.1748-1716.2011.02252.x 419

Adaptation to extreme environments in tardigrades Æ
N. Møbjerg et al. Acta Physiol 2011, 202, 409–420
2010. Transcriptome survey of the anhydrobiotic tardigrade
Milnesium tardigradum in comparison with Hypsibius
dujardini and Richtersius coronifer. BMC Genomics 11,
168.
May, R.M., Maria, M. & Guimard, J. 1964. Actions différentielles
des rayons x et ultraviolets sur le tardigrade Macrobiotus
areolatus, àl¢état actif et desséché. Bull Biol Fr Belg
98, 349–367.
Møbjerg, N. & Dahl, C. 1996. Studies on the morphology and
ultrastructure of the Malpighian tubules of Halobiotus
crispae Kristensen, 1982 (Eutardigrada). Zool J Linn Soc
116, 85–99.
Møbjerg, N., Jørgensen, A., Eibye-Jacobsen, J., Halberg, K.A.,
Persson, D. & Kristensen, R.M. 2007. New records on cyclomorphosis
in the marine eutardigrade Halobiotus crispae.
J Limnol 66(Suppl. 1), 132–140.
Nelson, D.R. 2001. Tardigrada. In: J. Thorp & A. Covich (eds)
Ecology and Classification of North American Freshwater
Invertebrates, 2nd edn, pp. 527–550. Academic Press, San
Diego.
Nelson, D.R. 2002. Current status of the Tardigrada: evolution
and ecology. Integr Comp Biol 42, 652–659.
Neumann, S., Reuner, A., Brümmer, F. & Schill, R.O. 2009.
DNA damage in storage cells of anhydrobiotic tardigrades.
Comp Biochem Physiol A Mol Integr Physiol 153, 425–
429.
Nielsen, C. 2001. Animal Evolution: Interrelationships of
the Living Phyla, 2nd edn. Oxford University Press,
Oxford.
Persson, D., Halberg, K.A., Jørgensen, A., Ricci, C., Møbjerg,
N. & Kristensen, R.M. 2011. Extreme stress tolerance in
tardigrades: surviving space conditions in low earth orbit.
J Zoolog Syst Evol Res: doi: 10.1111/j.1439-0469.2010.
00605.x.
Ramazzotti, G. 1962. Il Phylum Tardigrada. Mem Insti Ital
Idro Dott Marco De Marchi 14, 1–732.
Ramazzotti, G. 1972. Tardigradi delle isole Kerguelen e descrizione
della nouva specie Hypsibius (I) renaudi. Mem Ist
Ital Idrobiol 29, 141–144.
Ramazzotti, G. & Maucci, W. 1983. Il Phylum Tardigrada.
Terza edizione riveduta e corretta. Mem Insti Ital Idro Dott
Marco De Marchi 41, 1–1012.
Ramløv, H. & Westh, P. 1992. Survival of the cryptobiotic
tardigrade Adorybiotus coronifer during cooling to )196 °C:
effect of cooling rate, trehalose level and short term preacclimation.
Cryobiology 29, 125–130.
Ramløv, H. & Westh, P. 2001. Cryptobiosis in the eutardigrade
Adorybiotus (Richtersius) coronifer: tolerance to
alcohols, temperature and de novo protein synthesis. Zool
Anz 240, 517–523.
Rebecchi, L. & Bertolani, R. 1994. Maturative patterne of
ovary and testis in eutardigrades of fresh-water and terrestrial
habitats. Invert Rep Dev 26, 107–117.
Rebecchi, L., Altiero, T. & Guidetti, R. 2007. Anhydrobiosis:
the extreme limit of desiccation tolerance. Invertebr Survival
J 4, 65–81.
Rebecchi, L., Altiero, T., Guidetti, R., Cesari, M., Bertolani,
R., Negroni, M. & Rizzo, A.M. 2009a. Tardigrade resistance
to space effects: first results of experiments on the
LIFE-TARSE mission on FOTON-M3 (September 2007).
Astrobiology 6, 581–591.
Rebecchi, L., Cesari, M., Altiero, T., Frigieri, A. & Guidetti, R.
2009b. Survival and DNA degradation in anhydrobiotic
tardigrades. J Exp Biol 212, 4033–4039.
Renaud-Mornant, J. 1982. Species diversity in marine Tardigrada.
In: D.R. Nelson (ed.) Proceedings of the 3rd International
Symposium on Tardigrada, pp. 149–178. East
Tennessee State University Press, Johnson City.
Reuner, A., Hengherr, S., Brahim, M., Förster, F., Arndt, D.,
Reinhardt, R., Dandekar, T., Frohme, M., Brümmer, F. &
Schill, R.O. 2010. Stress response in tardigrades: differential
gene expression of molecular chaperones. Cell Stress Chaperones
15, 423–430.
Rizzo, A.M., Negroni, M., Altiero, T., Montorfano, G.,
Corsetto, P., Berselli, P., Berra, B., Guidetti, R. & Rebecchi,
L. 2010. Antioxidant defences in hydrated and desiccated
states of the tardigrade Paramacrobiotus richtersi. Comp
Biochem Physiol B 156, 115–121.
Sands, C.J., McInnes, S.J., Marley, N.J., Goodall-Copestake,
W.P., Convey, P. & Linse, K. 2008. Phylum Tardigrada: an
‘‘individual’’ approach. Cladistics 24, 1–11.
Schill, R.O. 2010. Anhydrobiotic abilities of tardigrades. In:
S. Hohmann (Series ed.) Topics in Current Genetics Vol. 21,
E. Lubzens, J. Cerda & M.S. Clark (eds) Dormancy and
resistance in harsh environments, pp. 133–146, Springer-
Verlag, Heidelberg.
Schill, R.O., Steinbrück, G.H.B. & Köhler, H.R. 2004. Stress
gene (hsp 70) and quantitative expression in Milnesium
tardigradum (Tardigrada) during active and cryptobiotic
stages. J Exp Biol 207, 1607–1613.
Schokraie, E., Hotz-Wagenblatt, A., Warnken, U., Mali, B.,
Frohme, M., Förster, F., Dandekar, T., Hengherr, S., Schill,
R.O. & Schnölzer, M. 2010. Proteomic analysis of tardigrades:
towards a better understanding of molecular mechanisms
by anhydrobiotic organisms. PLoS One 5, e9502.
Suzuki, A.C. 2003. Life history of Milnesium tardigradum
Doyère (Tardigrada) under a rearing environment. Zool Sci
20, 49–57.
Warner, A.H., Miroshnychenko, O., Kozarova, A., Vacratsis,
P.O., MacRae, T.H., Kim, J. & Clegg, J.S. 2010. Evidence
for multiple group 1 late embryogenesis abundant (LEA)
proteins in encysted embryos of Artemia and their organelles.
J Biochem 148, 581–592.
Westh, P. & Kristensen, R.M. 1992. Ice formation in the
freeze-tolerant eutardigrades Adorybiotus coronifer and
Amphibolus nebulosus studied by differential scanning calorimetry.
Polar Biol 12, 693–699.
Westh, P. & Ramløv, H. 1991. Trehalose accumulation in the
tardigrade Adorybiotus coronifer during anhydrobiosis.
J Exp Zool 258, 303–311.
Westh, P., Kristiansen, J. & Hvidt, A. 1991. Ice-nucleating
activity in the freezetolerant tardigrade Adorybiotus coronifer.
Comp Biochem Physiol A 99, 401–404.
Wright, J.C. 2001. Cryptobiosis 300 years on from van Leuwenhoek:
what have we learned about tardigrades Zool
Anz 240, 563–582.
Wright, J.C., Westh, P. & Ramløv, H. 1992. Cryptobiosis in
Tardigrada. Biol Rev 67, 1–29.
420
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Paper IV

Haugan et al. BMC Developmental Biology 2010, 10:56
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RESEARCH ARTICLE
Research article
Functional characterization of the vertebrate
primary ureter: Structure and ion transport
mechanisms of the pronephric duct in axolotl
larvae (Amphibia)
Birgitte M Haugan, Kenneth A Halberg, Åse Jespersen, Lea R Prehn and Nadja Møbjerg*
Open Access
Abstract
Background: Three kidney systems appear during vertebrate development: the pronephroi, mesonephroi and
metanephroi. The pronephric duct is the first or primary ureter of these kidney systems. Its role as a key player in the
induction of nephrogenic mesenchyme is well established. Here we investigate whether the duct is involved in urine
modification using larvae of the freshwater amphibian Ambystoma mexicanum (axolotl) as model.
Results: We investigated structural as well as physiological properties of the pronephric duct. The key elements of our
methodology were: using histology, light and transmission electron microscopy as well as confocal laser scanning
microscopy on fixed tissue and applying the microperfusion technique on isolated pronephric ducts in combination
with single cell microelectrode impalements. Our data show that the fully differentiated pronephric duct is composed
of a single layered epithelium consisting of one cell type comparable to the principal cell of the renal collecting duct
system. The cells are characterized by a prominent basolateral labyrinth and a relatively smooth apical surface with one
central cilium. Cellular impalements demonstrate the presence of apical Na + and K + conductances, as well as a large K +
conductance in the basolateral cell membrane. Immunolabeling experiments indicate heavy expression of Na + /K + -
ATPase in the basolateral labyrinth.
Conclusions: We propose that the pronephric duct is important for the subsequent modification of urine produced by
the pronephros. Our results indicate that it reabsorbs sodium and secretes potassium via channels present in the apical
cell membrane with the driving force for ion movement provided by the Na + /K + pump. This is to our knowledge the
first characterization of the pronephric duct, the precursor of the collecting duct system, which provides a model of
cell structure and basic mechanisms for ion transport. Such information may be important in understanding the
evolution of vertebrate kidney systems and human diseases associated with congenital malformations.
Background
During the development from embryo to adult life vertebrates
use a succession of kidney forms to maintain extracellular
fluid homeostasis and simultaneously rid the
body of nitrogenous wastes [1,2]. Three spatially and
temporally different kidney generations form from the
intermediate mesoderm in an anterior to posterior direction
i.e. the pronephroi, mesonephroi and metanephroi
[3]. The functional unit in these paired kidneys is the
* Correspondence: nmobjerg@bio.ku.dk
1 Department of Biology, University of Copenhagen, Universitetsparken, DK-
2100 Copenhagen, Denmark
Full list of author information is available at the end of the article
nephron, and it is composed of a filtration unit and a
renal tubule. Urine is produced by the filtration of blood
in the filtration unit, followed by the selective reabsorption
and secretion of ions, organic molecules and water
across highly specialized epithelia of the renal tubule [4].
Nephrons open into a ureter, and in the meso- and metanephros
they do so via a collecting duct system [5,6]. This
system is the site for the important final adjustment of
the urine. Vertebrate kidneys may produce urine, which is
either hypoosmotic (diluted), isoosmotic or hyperosmotic
(concentrated) relative to the body fluids [7-10].
This ability is a function of i) the evolutionary state of the
© 2010 Haugan et al; licensee BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons
Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in
any medium, provided the original work is properly cited.

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nephrons and ii) the regulation of filtration and of the
transport of inorganic ions, organic molecules and water
across the renal epithelia.
The first kidneys to form - the embryonic pronephric
kidneys - are the functional kidneys of fish and amphibian
larvae [11-16]. These are very simple kidneys composed
of a single nephron. A characteristic of vertebrate
kidney organogenesis is the development of a pronephric
duct in association with each pronephros [3]. These ducts
are the first or primary ureters of vertebrate kidney systems.
They form the collecting duct system of the mesoand
metanephroi, and they, and their derivates, are the
key players in the induction of the nephrogenic mesenchyme,
which forms these latter kidney generations. Few
functional studies exist on the pronephros and functional
studies of the duct are virtually lacking. Molecular studies
have been directed at mapping genes expressed in different
segments of the pronephric nephron, and several
recent reviews have highlighted the potential use of this
embryonic kidney in drug and human kidney disease
assessment [17-23]. To date, there has been little focus on
the role of the pronephric duct in urine modification and
it remains to be shown whether transepithelial transport
processes are present in this structure. Ultrastructural
investigations have shown duct cells with the characteristics
of an epithelium involved in active transport e.g.
many mitochondria and surface expansions of the basolateral
cell membranes [14,24]. In addition, gene expression
assays have indicated high expression of transporters
known to be involved in ion transport, such as the Na + /
K + -ATPase and the ROMK channel [17,18,25-27]. Collectively,
these data suggest that the pronephric duct may
play an important role in regulation of extracellular fluid
homeostasis. Therefore we ask the question: Is the duct
involved in urine modification
In amphibians the pronephros is a large organ, which is
functional for a considerable time, before it degenerates.
We investigate structural and functional characteristics of
the pronephric duct in the freshwater amphibian Ambystoma
mexicanum (axolotl). Members of the Ambystoma
genus have been used as models for the study of pronephric
structure, function, development and evolution
for more than a century [24,28,29] and the formation and
caudal migration of the pronephric duct in the axolotl has
been thoroughly investigated [30-32]. Numerous functional
studies exist on the mesonephric collecting duct
system of both urodele and anuran amphibians, which
provide detailed information on the transport characteristics
of these segments [Reviewed in [13]]. Our histological
examinations and dissections of axolotl larvae
indicate that the pronephros is functional from the time
of hatching to larval stage 54. We investigate duct morphology
and cellular transport mechanisms present in
larvae with functional pronephroi, and show that the primary
ureter is important for urine dilution in the axolotl.
The single cell type found in the ureter shows the characteristics
of the vertebrate collecting duct system principal
cell and our data indicate that it reabsorbs Na + and
secretes K + .
Results and Discussion
Identification of functional pronephroi and pronephric
ducts
We determined the interval in which the axolotl pronephros
and pronephric duct is functional by investigating
kidney structure in freshly dissected larvae and in
larvae prepared for histology (Figure 1 and 2). The pronephroi
of axolotl larvae are paired organs located on
each side of the dorsal aorta in the most anterior part of
the body cavity. They are visible from the outside on the
dorsal side of the larva as two small bulges behind the
gills (Figure 1A). Each of the two kidneys are composed of
a filtrating unit - a glomus originating from the dorsal
aorta, and a single convoluted renal tubule opening into
the coelom via two ciliated nephrostomes (Figure 1B).
The pronephros is fully functional when the axolotl larva
Figure 1 Axolotl larva stage 45 and 52. A. Dorsal view of recently
hatched axolotl larva (stage 45; forelimb present as limb buds). The
pronephroi can be seen as two small bulges behind the gills. B. Schematic
representation of the pronephric nephron in the stage 52 larva
as revealed from light- and transmission electron microscopy on serial
section of plastic embedded tissue. The pronephros consists of an external
glomus (gl) and a single renal tubule, which opens into the
coelom (co) via two ciliated nephrostomes (ne). In this late larval stage
the tubule is divided into two ciliated tubules (ci), two proximal tubule
branches (pt), a common proximal tubule, a ciliated intermediate segment
(is) and a distal tubule (dt). The distal tubule continues as the pronephric
duct (pd), which leaves the confines of the pronephros and
empties into the cloaca.

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A
pt
dt
nc
ne
gl
is
25 m
co
B
50 m
C
pt
mr
wd
mv
pd
co
bl
nu
mu
15m
5 m
Figure 2 Histology of pronephros and pronephric duct. A. Cross section of a stage 54 larva (forelimb completely developed) revealing the filtration
unit and the convoluted pronephric tubule. Araldite section, 1.5 μm, stained with toluidine blue. Blood is filtered in the external glomus (gl) and the
filtrate enters the coelom (co) before it is taken up into the renal tubule via ciliated nephrostomes (ne). In this late larval stage the tubule is characterized
by possessing a ciliated intermediate segment (is). nc, notochord; dt, distal tubule; pt, proximal tubule. B. Longitudinal section of pronephric duct
(stage 52 larva). Araldite section, 2 μm, stained with toluidine blue. The pronephric duct (pd) leaves the confines of the pronephros. co, coelom; mu,
muscle; pt, proximal tubule. INSERT: The Wolffian duct (wd) at the level of the caudal part of the mesonephros in a stage 54 larva. The duct epithelium
consists of two cell types: principal cells and intercalated, mitochondria-rich cells (mr). C. Transmission electron microscopy of pronephric duct shown
in figure 2B. The duct is composed of a single cell type characterized by a relative smooth apical surface with few microvilli (mv) and a well developed
basal labyrinth (bl) formed by the highly invaginated basal and to some extent lateral cell membranes. nu, nucleus.

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hatches from the egg at stage 44. This was confirmed by
the appearance of blood cells in the capillaries of the glomus,
which from embryonic stage 36 had a fully developed
endothelium and a visceral layer with podocytes.
From stage 44 the cilia of the nephrostomes points
toward the lumen of the renal tubule, indicating a passage
of fluid from the coelom. At this stage the kidney consists
of an external glomus and a renal tubule with the following
morphologically determined segments: two nephrostomes,
each connected to a branch of proximal tubule, a
common proximal tubule and a distal tubule. The pronephric
duct runs caudally as an extension of the distal
tubule opening into the cloaca. From the time of hatching
the kidney was observed to increase in overall size due to
further segmentation of the renal tubule i.e. the development
of a ciliated intermediary segment; present from
stage 52 (Figure 1B and 2A). The fully segmented pronephric
tubule consists of the following morphological
defined segments: two nephrostomial tubules, two proximal
tubule branches, a common proximal tubule, a ciliated
intermediary segment and a distal tubule (Figure
2A). At stage 52 the mesonephros was clearly visible and
functional as judged by glomerular maturation and presence
of blood cells in the mesonephric glomerular capillaries.
Hence, comparable to the situation found in
anuran amphibians [14,33] the two kidney generations
functionally overlap in axolotl larvae. At stage 54 the pronephros
reaches its maximal size. Gonadal primordia
were observed medioventral to the mesonephros (not
shown). These were undifferentiated and sex determination
was not possible. During the transition from stage 54
to latter stages, characterized by fully developed hind
limbs, overall pronephric tubule and glomus volume
decreased, marking pronephric degeneration. Pronephric
structure in stage 52-54 larvae of Ambystoma mexicanum
resembled the description by Christensen (1964) of the
functional pronephros in A. punctatum [24].
Pronephric duct structure
Structural examination of the pronephric duct in larvae
with functional pronephroi revealed that the duct consists
of a single cell type (Figure 2B and 2C). Thus, the
heterocellularity with intercalated mitochondria-rich
cells interposed between principal cells, characteristic of
the collecting duct system of latter kidney generations, is
not seen at this point (Figure 2B and 2C). This is comparable
to the situation in the green toad, Bufo viridis - a
terrestrial anuran amphibian [14]. The pronephric duct
cells in A. mexicanum are approximately 20 μm high in
early larval stages, but decrease in height to 10-15 μm in
stage 52 and 54 larvae (Figure 2C). They have a relatively
smooth apical surface. A single central cilium is present
(not shown) in addition to small and sparse microvilli,
which are more numerous at the point of the apical cell
junctions. The nucleus is regularly shaped, centrally
placed and contains a nucleolus and patches of heterochromatin.
There is a conspicuous basal labyrinth, and
lateral infoldings are seen as well (Figure 2C). The cytoplasm
contains many mitochondria in addition to a Golgi
complex and endoplasmatic reticulum. The morphology
of the duct changes at the level of the mesonephros in late
larval stages 52-54, revealing the presence of intercalated,
mitochondria-rich cells (Figure 2B, insert).
Ion transport mechanisms in the pronephric duct of larvae
stage 46-54
We examined if the pronephric duct participates in final
urine modification with the aid of glass microelectrodes
and ion substitution experiments in isolated and perfused
ducts dissected from 22 larvae. Figure 3A is a frequency
distribution of the membrane potential (V m ) of 64
impaled cells. The data show a broad distribution with an
Number of cells
V m (mV)
25
20
15
10
5
0
-20
-40
-60
-80
-100
A
0
-110 -90 -70 -50 -30
V m (mV)
B
high K
2 min
V m (mV)
-40
-60
-80
-100
high K
Figure 3 Voltage recordings from single cells in isolated and perfused
ducts. A. Frequency distribution of the membrane potential
(V m ) in 64 cells from pronephric ducts dissected from axolotl larvae in
the stage 46-54. B. Original voltage trace from single cell. Effect of raising
bath [K + ] from 3 to 20 mmol/l. V m depolarized indicating the presence
of a basolateral K + conductance. C. Summary data illustrating the
effect on V m of the bath K + concentration step (n = 29).
0
-20
C
-10

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average V m between -75 and -80 mV. Transport characteristics
of the duct did not seem to differ between early
and late larval stages. As shown in figure 3B and 3C, raising
the K + concentration in the basal solution from 3 to
20 mmol/l resulted in a large, reversible depolarization of
V m , revealing the presence of a large basolateral K + conductance.
This indicates the presence of K + channels in
the basolateral cell membrane. We examined whether the
duct has luminal electrogenic transporters or channels in
experiments with luminal K + and Na + steps. Figure 4
V m (mV)
0
-20
-40
-60
-80
A
low Na
high K
low Na
shows the result of these substitution experiments. V m
hyperpolarized upon a decrease in luminal Na + concentration
from 102 to 7 mmol/l and depolarized upon an
increase in luminal K + concentration from 3 to 20 mmol/
l. This indicates that the luminal (apical) cell membrane
possesses Na + as well as K + channels. In order to identify
an ion pump, which can provide the driving force for
luminal uptake of Na + as well as K + secretion, we isolated
pronephric ducts and performed immunolabeling with
an antibody directed against the Na + -K + -ATPase α-subunit.
As shown in figure 5, the Na + -K + -ATPase is highly
expressed in the pronephric ducts from these larvae, and
is entirely localized to the lateral and highly invaginated
basolateral cell membranes.
Electrophysiological studies performed on the mesonephric
collecting duct system of amphibians, have indicated
that principal cells in aquatic urodeles, have a large
apical Na + conductance and no, or very small, K + conductance.
However, in terrestrial anurans, K + secretion
through apical K + channels seems a major task of the
principal cells [34-39]. In the current study we provide
evidence for a principal cell, which has the characteristics
of the mammalian collecting duct principal cell, i.e. with
luminal Na + as well as K + conductances.
-100
5 min
A
0
B
-20 low Na -20 high K
0
C
V m (mV)
-40
-60
V m (mV)
-40
-60
B
25ìm
-80
-80
-100
-100
Figure 4 Electrophysiological response to luminal fluid exchange.
A. Original voltage trace from single cell. Effect of changing
luminal [Na + ] from 102 to 7 mmol/l and luminal [K + ] from 3 to 20
mmol/l. V m hyperpolarized upon the decrease in luminal [Na + ] and depolarized
upon an increase in luminal [K + ]. This indicates that the luminal
(apical) cell membrane possesses Na + as well as K + conductances.
B. Summary data illustrating the effect on V m of concentration steps in
luminal Na + (n = 6) and K + (n = 5).
DAPI
75ìm + +
Na -K -ATPase
Figure 5 Na + -K + -ATPase expression. A. The Na + -K + -ATPase is highly
expressed in the pronephric ducts and entirely localized to the lateral
and highly invaginated basolateral cell membranes. B. Na + -K + -ATPase
expression in duct counterstained with DAPI. Images are three-dimensional
reconstructions of the original CLSM z-series, showing a median
longitudinal section of the pronephric duct.

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Conclusions
We show that the pronephric duct, which is the first or
primary ureter in all vertebrates, participates in urine
adjustment in the axolotl. The cells constituting the duct
are on the ultrastructural as well as cell physiological level
comparable to principal cells found in the first segments
of the mammalian collecting duct system [5,6,14,40-42].
Notably, the pronephric duct lacks intercalated, mitochondria-rich
cells. We propose that the duct is important
for urine dilution through NaCl reabsorbtion, and
that it in addition participates in the regulation of K +
homeostasis. Figure 6 provides a model of the ion transport
mechanisms, which we suggest are present in the
duct cell. In this model a Na + -K + -ATPase in the basolateral
cell membrane pumps Na + out of the cell and thereby
provides the driving force for apical uptake of Na +
through channels. The epithelial sodium channel (ENaC)
is a likely candidate mediating this apical Na + uptake [43-
45]. Na + exits the cell through the pump. K + is secreted
through apical channels, and is recycled for the pump
across the basolateral cell membrane. It is highly probable
that ROMK channels, known to mediate K + secretion in
the mammalian collecting duct system [42], and shown to
be expressed in amphibian pronephric ducts [18], mediate
K + secretion across the apical cell membrane. The
active transport of Na + would create a lumen-negative
transepithelial potential, and Cl - would presumably follow
passively through the paracellular pathway of this epithelium
[45]. This is to our knowledge the first characterization
of the pronephric duct-the precursor of the
+
Na
+
Na
+
+
K K
ATP
Figure 6 Suggested model for ion transport mechanisms in the
cells of the vertebrate primary ureter. A Na + -K + -ATPase in the basolateral
cell membrane pumps Na + out of the cell and thereby provides
the driving force for apical uptake of Na + through channels. K + is secreted
across the apical cell membrane through channels and recycled for
the pump across the basolateral cell membrane.
collecting duct system- which provides detailed information
on cell structure and the basic mechanisms for ion
transport.
Methods
Animals
Specimens of the Mexican axolotl Ambystoma mexicanum
(Shaw and Nodder, 1798) came from the animal stable
of the August Krogh Building, part of the Campus
Animal Research Facility at University of Copenhagen.
Staging were performed according to [46] for embryos
and the larvae were designated according to the degree of
limb development as defined by the Ambystoma Genetic
Stock Center, University of Kentucky; http://www.ambystoma.org.
Larvae used for experiments, were in the stage
44 to 54. They were euthanized by decapitation, followed
by brain destruction, and were subsequently either prepared
for histology or pronephric ducts (o.d. 50-70 μm,
dissected length 300-1000 μm) were free hand dissected
at 6°C and prepared for microperfusion experiments or
immunolabeling. The pronephric ducts were dissected
from the region in front of the mesonephros. Dissections
were performed in media containing (in mmol/l): 75
NaCl, 20 NaHCO 3 , 3.0 KCl, 1.8 CaCl 2 , 1.0 MgSO 4 , 0.8
Na 2 HPO 4 , 0.2 NaH 2 PO 4 , 5.5 glucose, 3.3 glycine, 0.4 PVP,
5.0 HEPES, titrated to pH 7.8 with NaOH.
Histology, Light and Transmission Electron Microscopy
Dissections were performed for every developmental
stage of post-hatched axolotls from stage 44 to 57. In
addition, we examined pronephric development in
embryos. Light microscopic imaging of live and dissected
larvae was performed using Zeiss Stemi 2000-CS and
Leica MZ 16 microscopes equipped with an Infinity X
Digital Camera (DeltaPix, Denmark). For histology, a
total of 31 larvae in the stage 44-54 and 19 embryos from
stage 21 to 44 were used. Specimens were fixed for 12
hours at room temperature in an aldehyde fixative containing:
1.2% glutaraldehyde, 1% paraformaldehyde, 0.05
mol/l sucrose and 0.05 mol/l sodium cacodylate buffer
(pH 7.4) and subsequently rinsed and stored in 0.05 mol/l
sodium cacodylate buffer with 0.05 mol/l sucrose. Following
1 hour's post fixation in 2% OsO 4 with 0.1 mol/l
sodium cacodylate, specimens were dehydrated through a
graded series of ethanol and propylenoxide and embedded
in Araldite. For light microscopy, 1.5 μm sections
were cut with glass knives on a Leica ultramicrotome EM
UC6 and stained with toluidine blue. Ultrathin sections
for transmission electron microscopy were cut on the
same microtome with a Diatome diamond knife and subsequently
stained with uranyl acetate and lead citrate.
Transmission electron microscopic images were acquired
using JEOL 100SX and JEOL JEM 1011 transmission electron
microscopes. Kodak negatives obtained from the

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JEOL 100SX were digitized using an Epson Perfection
4990 Photo scanner. The JEOL JEM-1011 was equipped
with a GATAN digital camera. Digital images were optimized
for contrast and color using CorelDraw X4.
Microperfusion and cellular impalements
Pronephric ducts were transferred to a bath chamber
mounted on an inverted microscope and perfused in vitro
with a set of pipettes made to fit the diameter of the
tubules [37-39]. The tubule perfusion system used (Luigs
& Neumann, Germany) was designed for accurate adjustment
and movement of concentric pipettes [47]. Holding
and perfusion pipettes were hand made from glass tubing
(Drummond Scientific Company, PA, USA; holding
pipettes: o.d. 2.1 mm, i.d. 1.6 mm; perfusion pipettes: o.d.
1.2 mm, i.d. 1.0 mm) in a microforge equipped with a
microscope (SM II/1 Puller from Luigs & Neumann, Germany).
The tubules were perfused with a single-barrelled
perfusion pipette containing a small glass capillary (o.d.
0.3 mm, i.d. 0.2 mm, Drummond Scientific Company, PA,
USA) connected to a manual Hamilton valve (Hamilton
Co., NV, USA) with a four-way distribution system. Fluid
exchange during luminal perfusion experiments was
made through this capillary, the tip of the capillary being
placed close to the opening of the perfusion pipette,
ensuring fast fluid exchange in the tubule. Luminal perfusion
was performed either by hand through a syringe
connected to one of the ports in the valve or it was gravity-driven
through a port connected to a fluid filled reservoir.
The pressure applied was adjusted by monitoring
tubule diameter, ensuring that the tubule neither collapsed
nor over expanded. The bath was perfused at 8 ml/
min and fluid exchange was performed through beakers
attached to the outside of the Faraday cage, which surrounds
the microperfusion set-up.
Perfusions of tubule bath and luminal fluid were carried
out at room temperature with a perfusion solution containing
(in mmol/l): 75 NaCl, 25 NaHCO 3 , 3.0 KCl, 1.8
CaCl 2 , 1.0 MgSO 4 , 0.8 Na 2 HPO 4 , 0.2 NaH 2 PO 4 . The perfusion
solution was equilibrated with 1.8% CO 2 in O 2 and
had a measured pH of 7.8. Experimental solutions with
different sodium and potassium concentrations were prepared
from this control solution. In the high K + solution,
the K + concentration was raised to 20 mmol/l by equimolar
substitution with Na + . In the low Na + solution with a
[Na + ] of 6.8 mmol/l, Na + was replaced by choline or N-
methyl-D-glucamine (NMDG + ) titrated with HCl.
Pronephric duct cells were impaled across the basal cell
membrane with KCl (1-3 mol/l) filled glass microelectrodes
(R electrode ≈ 100 MΩ) and the membrane potential
(V m ) was recorded with respect to the grounded bath.
The microelectrodes were pulled from borosilicate glass
with filament (Clark Electromedical, UK) on a vertical
electrode puller (Narishige, Japan). Impalements were
achieved by placing the microelectrode tip against the
basal surface of the cell and gently taping the micromanipulator
(Leitz, Germany) on which the electrodes were
mounted. Voltage recordings were made by a WPI Duo
773 electrometer (World Precision Instruments, USA)
and digitized by a PowerLab/4S data acquisition system
(ADInstruments, Australia). The recording of V m was
accepted if the impalement was achieved by a sudden
change in the potential read by the electrode and if the
impalement was stable.
The results are based on 64 cell impalements made in
22 pronephric ducts dissected from 22 axolotl larvae in
the stage 46-54. Figures were made in Origin 7.5 (Microcal,
USA) and CorelDRAW X4.
Immunolabeling, Confocal Laser Scanning Microscopy and
3D reconstruction
For identification and localization of the Na + -K + -ATPase,
three separate immunolabeling experiments were conducted
with equal results covering larvae in stages 46-54.
In each experiment, pronephric ducts were isolated from
four to five specimens and subsequently fixed on ice for
approximately 60 minutes in 3% paraformaldehyde buffered
to pH 7.4 with 0.1 mol/l sodium cacodylate. After
being rinsed in PBS (perfusion solution), the tissue was
incubated overnight at 4°C in PBS containing 10% goat
serum (Invitrogen, CA, USA), 1% triton-X and the Na + -
K + ATPase monoclonal mouse antibody α5-IgG (10 μg/
ml). The α5 antibody developed by D.M. Fambrough was
obtained from the Developmental Studies Hybridoma
Bank developed under the auspices of the NICHD and
maintained by The University of Iowa, Department of
Biology (Iowa City, IA 52242). Following an extensive
wash in PBS, the pronephric ducts were incubated with
Alexa-488-conjugated goat anti-mouse IgG (1:100, Invitrogen,
CA, USA) overnight at 4°C. Following rinses in
PBS, the tissue was mounted on glass coverslips in Vecta
shield (Vector Laboratories Inc., CA, USA). In some
preparations, the renal tubules were incubated in DAPI
(1:250, Invitrogen, CA, USA) for approximately 5 min
and washed in PBS, prior to mounting. Image acquisition
was performed on a Leica DM RXE 6 TL inverted microscope
equipped with a Leica TCS SP2 AOBS confocal
laser scanning unit, using the 488 nm line of an argon/
crypton laser. The image series were processed and
edited in the 3-D reconstruction software IMARIS (Bitplane
AG, Zürich, Switzerland). The confocal images are
based on 150-170 optical sections of a Z-series performed
at intervals of 0.3-0.5 μm. All control preparations
were negative for immunostaining.
Abbreviations
bl: basal labyrinth; ci: ciliated tubule; co: coelom; dt: distal tubule; gl: glomus; is:
ciliated intermediate segment; nc: notochord; ne: nephrostome; nu: nucleus;

Haugan et al. BMC Developmental Biology 2010, 10:56
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mr: intercalated mitochondria-rich cell; mu: muscle; mv: microvilli; pd: pronephric
duct; pt: proximal tubule; V m : membrane potential; wd: Wolffian duct.
Authors' contributions
NM conceived and designed the study. BMH and NM fixed larvae and performed
the dissections for the structural investigation. BMH sectioned specimens,
and made LM and TEM investigations with help from ÅJ and NM. NM
made the microperfusion experiments and microelectrode impalements. KAH,
LRP and NM carried out immunostaining experiments and KAH performed
CLSM and prepared the 3D images. All authors participated in discussions and
interpretation of the data. NM wrote the paper with inputs from the other
authors. All authors read and approved the final version of the manuscript.
Acknowledgements
We sincerely thank Mrs. Jette Lyby Michelsen and Mrs. Kristine J.K. Sørensen for
technical assistance. Funding came from the 2008 Faculty of Science, University
of Copenhagen Freja-Programme and from the Carlsberg Foundation
(grant numbers: 2004_04_0572; 2006_01_0534; 2008_01_0466). The funders
had no role in study design, data collection and analysis, decision to publish, or
preparation of the manuscript.
Author Details
Department of Biology, University of Copenhagen, Universitetsparken, DK-
2100 Copenhagen, Denmark
Received: 18 September 2009 Accepted: 27 May 2010
Published: 27 May 2010
© This BMC 2010 is article Developmental an Haugan Open is available Access et al; Biology licensee from: article 2010, http://www.biomedcentral.com/1471-213X/10/56
distributed BioMed 10:56 Central under Ltd. the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
References
1. Evans DH: Osmotic and ionic regulation: Cells and animals Boca Raton: CRC
Press; 2009.
2. Vize PD, Woolf AS, Bard JBL: The Kidney: From normal development to
congenital disease London: Academic Press; 2003.
3. Saxén L: Organogenesis of the kidney Cambridge: Cambridge University
Press; 1987.
4. Dantzler WH: Comparative aspects of renal function. In The Kidney:
Physiology and Pathophysiology 2nd edition. Edited by: Seldin D, Giebisch
G. New York: Raven Press; 1992:885-942.
5. Møbjerg N, Jespersen A, Wilkinson M: Morphology of the kidney in the
West African caecilian, Geotrypetes seraphini (Amphibia, Gymnophiona,
Caeciliidae). Journal of Morphology 2004, 262:583-607.
6. Kriz W, Kaissling B: Structural and functional organization of the
mammalian kidney. In Seldin and Giebisch's the kidney. Physiology and
pathophysiology Edited by: Alpern RJ, Herbert SC. Burlington, MA:
Academic Press; 2008:479-563.
7. Greger R: Principles of renal transport; concentration and dilution of
urine. In Comprehensive human physiology. From cellular mechanisms to
integration Volume 2. Edited by: Greger R, Windhorst U. Berlin Heidelberg:
Springer; 1996:1489-1516.
8. Giebisch G, Windhager E: Urine concentration and dilution. In Medical
physiology. A cellular and molecular approach Edited by: Boron WF,
Boulpaep EL. Philadelphia: Saunders; 2005:828-844.
9. Larsen EH, Møbjerg N, Nielsen R: Application of the Na + recirculation
theory to ion coupled water transport in low- and high resistance
osmoregulatory epithelia. Comparative Biochemistry and Physiology
2007, 148:101-116.
10. Larsen EH, Møbjerg N: Na + recirculation and isosmotic transport.
Journal of Membrane Biology 2006, 212:1-15.
11. Brandli AW: Towards a molecular anatomy of the Xenopus pronephric
kidney. International Journal of Developmental Biology 1999, 43:381-395.
12. Drummond IA, Majumdar A, Hentschel H, Elger M, Solnica-Krezel L, Schier
AF, Neuhauss SCF, Stemple DL, Zwartkruis F, Rangini Z, et al.: Early
development of the zebrafish pronephros and analysis of mutations
affecting pronephric function. Development 1998, 125:4655-4667.
13. Hillyard SD, Møbjerg N, Tanaka S, Larsen EH: Osmotic and ion regulation
in amphibians. In Osmotic and ionic regulation: Cells and animals 1st
edition. Edited by: Evans DH. Boca Raton: CRC Press; 2009:367-441.
14. Møbjerg N, Larsen EH, Jespersen A: Morphology of the kidney in larvae
of Bufo viridis (Amphibia, Anura, Bufonidae). Journal of Morphology
2000, 245:177-195.
15. Vize PD: Embryonic kidneys and other nephrogenic models. In The
Kidney: From normal development to congenital disease Edited by: Vize PD,
Woolf AS, Bard JBL. London: Academic Press; 2003:1-6.
16. Vize PD, Seufert DW, Carroll TJ, Wallingford JB: Model systems for the
study of kidney development: Use of the pronephros in the analysis of
organ induction and patterning. Developmental Biology 1997,
188:189-204.
17. Zhou XL, Vize PD: Proximo-distal specialization of epithelial transport
processes within the Xenopus pronephric kidney tubules.
Developmental Biology 2004, 271:322-338.
18. Tran U, Pickney LM, Ozpolat BD, Wessely O: Xenopus Bicaudal-C is
required for the differentiation of the amphibian pronephros.
Developmental Biology 2007, 307:152-164.
19. Raciti D, Reggiani L, Geffers L, Jiang Q, Bacchion F, Subrizi AE, Clements D,
Tindal C, Davidson DR, Kaissling B, et al.: Organization of the pronephric
kidney revealed by large-scale gene expression mapping. Genome
Biology 2008, 9:.
20. Wingert RA, Davidson AJ: The zebrafish pronephros: A model to study
nephron segmentation. Kidney International 2008, 73:1120-1127.
21. Vasilyev A, Liu Y, Mudumana S, Mangos S, Lam PY, Majumdar A, Zhao JH,
Poon KL, Kondrychyn I, Korzh V, et al.: Collective cell migration drives
morphogenesis of the kidney nephron. Plos Biology 2009, 7:101-114.
22. Wheeler GN, Brandli AW: Simple vertebrate models for chemical
genetics and drug discovery screens: Lessons from Zebrafish and
Xenopus. Developmental Dynamics 2009, 238:1287-1308.
23. Vize PD, Carroll TJ, Wallingford JB: Induction, development and
physiology of the pronephric tubules. In The Kidney: From normal
development to congenital disease Edited by: Vize PD, Woolf AS, Bard JBL.
London: Academic Press; 2003:19-50.
24. Christensen AK: Structure of functional pronephros in larvae of
Ambystoma opacum as studied by light and electron microscopy.
American Journal of Anatomy 1964, 115:257-278.
25. Eid SR, Brandli AW: Xenopus Na, K-ATPase: primary sequence of the beta
2 subunit and in situ localization of alpha 1, beta 1, and gamma
expression during pronephric kidney development. Differentiation
2001, 68:115-125.
26. Kumano T, Konno N, Wakasugi T, Matsuda K, Yoshizawa H, Uchiyama M:
Cellular localization of a putative Na + /H + exchanger 3 during ontogeny
in the pronephros and mesonephros of the Japanese black
salamander (Hynobius nigrescens Stejneger). Cell and Tissue Research
2008, 331:675-685.
27. Uochi T, Takahashi S, Ninomiya H, Fukui A, Asashima M: The Na + , K + -
ATPase alpha subunit requires gastrulation in the Xenopus embryo.
Development Growth & Differentiation 1997, 39:571-580.
28. Field HH: The development of the pronephros and segmental duct in
Amphibia Cambridge, USA: Bulletin of the Museum of Comparative
Zoology; 1891.
29. Howland RB: On the effect of removal of the pronephros of the
amphibian embryo. Proceedings of the National Academy of Sciences of
the United States of America 1916, 2:231-234.
30. Drawbridge J, Steinberg MS: Morphogenesis of the axolotl pronephric
duct: A model system for the study of cell migration in vivo.
International Journal of Developmental Biology 1996, 40:709-713.
31. Gillespie LL, Armstrong JB: Formation of the pronephros and pronephric
duct rudiment in the Mexican axolotl. Journal of Morphology 1985,
185:217-222.
32. Poole TJ, Steinberg MS: Amphibian pronephric duct morphogenesis -
segregation, cell rearrangement and directed migration of the
Ambystoma duct rudiment. Journal of Embryology and Experimental
Morphology 1981, 63:1-16.
33. Jaffee OC: Morphogenesis of the pronephros of the leopard frog (Rana
pipiens). Journal of Morphology 1954, 95:109-123.
34. Dietl P, Stanton BA: The amphibian distal nephron. In New insights in
vertebrate kidney function Edited by: Brown JA, Balment RJ, Rankin JC.
Cambridge: Cambridge University Press; 1993:115-134.
35. Horisberger J, Hunter M, Stanton B, Giebisch G: The collecting tubule of
Amphiuma II. Effects of potassium adaptation. American Journal of
Physiology 1987, 253:F1273-F1282.
36. Hunter M, Horisberger J-D, Stanton B, Giebisch G: The collecting tubule
of Amphiuma I. Electrophysiological characterization. American Journal
of Physiology 1987, 253:F1263-F1272.

Haugan et al. BMC Developmental Biology 2010, 10:56
http://www.biomedcentral.com/1471-213X/10/56
Page 9 of 9
37. Møbjerg N, Larsen EH, Novak I: K + transport in the mesonephric
collecting duct system of the toad Bufo bufo: microelectrode
recordings from isolated and perfused tubules. Journal of Experimental
Biology 2002, 205:897-904.
38. Møbjerg N, Larsen EH, Novak I: Ion transport mechanisms in the
mesonephric collecting duct system of the toad Bufo bufo:
microelectrode recordings from isolated and perfused tubules.
Comparative Biochemistry and Physiology A-Molecular & Integrative
Physiology 2004, 137:585-595.
39. Møbjerg N, Werner A, Hansen SM, Novak I: Physiological and molecular
mechanisms of inorganic phosphate handling in the toad Bufo bufo.
Pflugers Archiv - European Journal of Physiology 2007, 454:101-113.
40. Koeppen BM: Conductive properties of the rabbit outer medullary
collecting duct: outer stripe. American Journal of Physiology - Renal
Physiology 1986, 250:F70-F76.
41. Schlatter E: Regulation of ion channels in the cortical collecting duct.
Renal Physiology and Biochemistry 1993, 16:21-36.
42. Wang WH, Giebisch G: Regulation of potassium (K) handling in the renal
collecting duct. Pflugers Archiv-European Journal of Physiology 2009,
458:157-168.
43. Garty H, Palmer LG: Epithelial sodium channels: Function, structure, and
regulation. Physiological Reviews 1997, 77:359-396.
44. Loffing J, Korbmacher C: Regulated sodium transport in the renal
connecting tubule (CNT) via the epithelial sodium channel (ENaC).
Pflugers Archiv-European Journal of Physiology 2009, 458:111-135.
45. Schlatter E, Greger R, Schafer JA: Principal cells of cortical collecting
ducts of the rat are not a route of transepithelial Cl - transport. Pflugers
Archiv-European Journal of Physiology 1990, 417:317-323.
46. Bordzilovskaya NP, Dettlaf TA, Duhan ST, Malacinski GM: Developmentalstage
series of axolotl embryos. In Developmental Biology of the Axolotl
Edited by: Armstrong JB, Malacinski GM. New York: Oxford University
Press; 1989:201-219.
47. Greger R, Hampel W: A modified system for in vitro perfusion of isolated
renal tubules. Pflugers Archiv - European Journal of Physiology 1981,
389:175-176.
doi: 10.1186/1471-213X-10-56
Cite this article as: Haugan et al., Functional characterization of the vertebrate
primary ureter: Structure and ion transport mechanisms of the pronephric
duct in axolotl larvae (Amphibia) BMC Developmental Biology 2010,
10:56

Paper V

2803
The Journal of Experimental Biology 212, 2803-2811
Published by The Company of Biologists 2009
doi:10.1242/jeb.029413
Cyclomorphosis in Tardigrada: adaptation to environmental constraints
Kenneth Agerlin Halberg 1 , Dennis Persson 1,2 , Hans Ramløv 3 , Peter Westh 3 , Reinhardt Møbjerg Kristensen 2
and Nadja Møbjerg 1, *
1 Department of Biology, University of Copenhagen, August Krogh Building, Universitetsparken 13, DK-2100 Copenhagen Ø,
Denmark, 2 Natural History Museum of Denmark, Zoological Museum, Invertebrate Department, Universitetsparken 15, DK-2100
Copenhagen Ø, Denmark and 3 Department of Nature, Systems and Models, University of Roskilde, Universitetsvej 1, DK-4000
Roskilde, Denmark
*Author for correspondence (e-mail: nmobjerg@bio.ku.dk)
Accepted 9 June 2009
SUMMARY
Tardigrades exhibit a remarkable resilience against environmental extremes. In the present study, we investigate mechanisms of
survival and physiological adaptations associated with sub-zero temperatures and severe osmotic stress in two commonly found
cyclomorphic stages of the marine eutardigrade Halobiotus crispae. Our results show that only animals in the so-called
pseudosimplex 1 stage are freeze tolerant. In pseudosimplex 1, as well as active-stage animals kept at a salinity of 20 ppt, ice
formation proceeds rapidly at a crystallization temperature of around –20°C, revealing extensive supercooling in both stages,
while excluding the presence of physiologically relevant ice-nucleating agents. Experiments on osmotic stress tolerance show
that the active stage tolerates the largest range of salinities. Changes in body volume and hemolymph osmolality of active-stage
specimens (350–500 μm) were measured following salinity transfers from 20 ppt. Hemolymph osmolality at 20 ppt was
approximately 950 mOsm kg –1 . Exposure to hypo-osmotic stress in 2 and 10 ppt caused (1) rapid swelling followed by a regulatory
volume decrease, with body volume reaching control levels after 48 h and (2) decrease in hemolymph osmolality followed by a
stabilization at significantly lower osmolalities. Exposure to hyperosmotic stress in 40 ppt caused (1) rapid volume reduction,
followed by a regulatory increase, but with a new steady-state after 24 h below control values and (2) significant increase in
hemolymph osmolality. At any investigated external salinity, active-stage H. crispae hyper-regulate, indicating a high water
turnover and excretion of dilute urine. This is likely a general feature of eutardigrades.
Key words: cyclomorphosis, environmental stress, freeze tolerance, Halobiotus crispae, invertebrate, osmoregulation, tardigrade, volume
regulation.
INTRODUCTION
The phylum Tardigrada comprises a group of hydrophilous micrometazoans,
exhibiting close affinities to the euarthropod complex
(Garey et al., 1996; Giribet et al., 1996; Mallatt et al., 2004). They
occupy a range of niches in terrestrial, freshwater and marine
environments from continental Antarctica (Convey and McInnes,
2005) to the icecap of Greenland (Grøngaard et al., 1999) yet are
especially abundant in mosses and lichens, where they constitute a
major component of the cryptic fauna. Along with nematodes and
rotifers, selected species of tardigrades exhibit a remarkable
resilience against physical extremes, including low and high
temperatures (–253°C to +151°C), ionizing radiation (up to
6000Gy), vacuum, high pressure (up to 600MPa) and extreme
desiccation (Ramløv and Westh, 1992; Westh and Kristensen, 1992;
Ramløv and Westh, 2001; Schill et al., 2004; Horikawa et al., 2006;
Jönson and Schill, 2007; Hengherr et al., 2008; Hengherr et al.,
2009). However, the underlying physiological and biochemical
mechanisms mediating these unique tolerances are still largely
unidentified and represent an exciting challenge to contemporary
biology.
The marine eutardigrade Halobiotus crispae Kristensen 1982
(Fig.1) colonizes tidal and subtidal habitats at numerous localities
throughout the northern hemisphere (Møbjerg et al., 2007). This
species is characterized by the appearance of seasonal cyclic
changes in morphology, i.e. cyclomorphosis (Kristensen, 1982).
Three distinct cyclomorphic stages have been recognized: (1) the
active stage, (2) the pseudosimplex 1 (P1) stage and (3) the
pseudosimplex 2 (P2) stage (Møbjerg et al., 2007). The defining
physiological and biochemical characteristics of the individual stages
are largely unknown but most likely correlate with dominant abiotic
factors. A ubiquitous factor in all tidal and subtidal habitats is the
large temporal and spatial fluctuations in external salinity. Yet,
additional adaptations are necessary at high latitudes to ensure winter
survival due to prolonged exposure to subzero temperatures. In the
present study, we focus on the adaptive significance of the two main
cyclomorphic stages in H. crispae, i.e. the active stage corresponding
to the reproductive stage of other tardigrades and the P1 stage, a
hibernation stage, which is comparable to the cysts found in other
tardigrades (e.g. Guidetti et al., 2008). We do not deal with the P2
stage, which is a sexual maturation stage that has not yet been
reported from other tardigrades. Our preliminary data, however,
suggest that this stage has a unique osmoregulatory profile. We show
that the transition between the active and P1 stages is associated
with profound changes in the physiology of the animal. The P1 stage
is the only stage at which H. crispae survives internal ice formation.
The active stage tolerates large shifts in ambient salinity and we
investigate in detail the volume and osmoregulatory capacity of this
stage. Our study presents the first detailed analysis of osmoregulation
in tardigrades. The data show that active-stage H. crispae hyperregulate
at any investigated external salinity, which would indicate
excretion of dilute urine. This is likely to be a general feature of
eutardigrades, which all possess Malpighian tubules.
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K. A. Halberg and others
Fig. 1. SEM investigation of Halobiotus crispae from Vellerup Vig, Denmark. (A) Overview of P1 stage indicating the areas shown in B and C. The thick outer
cuticle functionally isolates the animal from the surroundings (scale bar=100 μm). (B) Close-up of the head region of P1. Notice that the mouth is closed by
cuticular thickenings (scale bar=25 μm). (C) Close-up of the posterior area of the P1 stage. As shown for the mouth, the cloaca is closed (scale bar=10 μm).
(D) Close-up of the head region of the active stage. Note the six peribuccal sensory organs (*) that surround the open mouth (scale bar=25 μm). (E) Close-up
of the posterior area of the active stage, revealing the open tri-lobed cloaca (scale bar=10 μm).
MATERIALS AND METHODS
Tardigrade sampling
Specimens of Halobiotus crispae were collected at regular intervals
in the period 2005 to 2008 at Vellerup Vig, Isefjord, Denmark
(55°44.209N, 11°51.365E) and at Nipisat Bay, Disko Island, West
Greenland (69°25.934N, 54°10.768E) in August 2006. At Vellerup
Vig, bottom samples were collected at an approximate depth of
1.0–2.5m, while samples from Nipisat Bay were taken in the subtidal
zone 3–4cm below low tide. With the exception of the data on
osmotic stress tolerance presented in Fig.3, the obtained results are
based entirely on animals collected at Vellerup Vig. Detailed
descriptions of the two localities can be found elsewhere (Kristensen,
1982; Møbjerg et al., 2007). Collected samples were freshwatershocked,
decanted into a conical net (mesh size 62 μm) and
transferred to Petri dishes. These dishes were supplied with fresh
seawater (SW; 18–20ppt; pH8–9) and substrate from the locality.
Tardigrades were localized using a Leica MZ16 stereomicroscope
(Leica Microsystems, Wetzlar, Germany), and primarily found on
the haptera of various filamentous algae present in the substrate.
Isolated tardigrades from Vellerup Vig were kept at 4°C in SW for
periods of up to 6months by regularly supplying fresh substrate
from the locality. Different cyclomorphic stages (see Fig.1) were
identified using an Olympus BX 51 interference-contrast microscope
(Olympus, Tokyo, Japan).
Scanning electron microscopy
For scanning electron microscopy, specimens were fixed in 2.5%
glutaraldehyde in 0.1moll –1 sodium cacodylate buffer (pH7.4),
rinsed in the buffer and subsequently postfixed in 1% OsO 4 in
0.1moll –1 sodium cacodylate buffer (pH7.4). Following fixation,
the specimens were dehydrated through a graded series of ethanol
and acetone. They were critical point dried (Bal-Tec CPD 030 critical
point dryer, Bal-Tec Union, Balzers, Liechtenstein), mounted on
aluminum stubs, sputter-coated with platinum–palladium (thickness
~12nm) using a JEOL JFC-2300HR (JEOL, Tokyo, Japan) and
examined in a JEOL JSM-6335F Field Emission scanning electron
microscope (JEOL, Japan).
Cold hardiness
Six groups of 10 animals in both the active and P1 stage were
transferred to Eppendorf tubes containing 1.5ml of SW from
Vellerup Vig (20ppt). The samples were cooled to a constant
temperature of –20°C at a cooling rate of approximately 1°Cmin –1
(Block, 1991) and held at the target temperature for a period of 24h.
The animals were thawed at room temperature and the survival
assessed successively over the course of 96h. Animals retaining
locomotory function or responsive to tactile stimuli following this
period were considered alive.
As subzero temperatures may be experienced for longer periods
of time in Arctic habitats, the long-term survival at subzero
temperatures was investigated. An additional six groups of 10
specimens in each cyclomorphic stage were frozen to –20°C at a
cooling rate of approximately 1°Cmin –1 and kept frozen for a total
of 36days. Survival was assessed as described above.
Differential scanning calorimetry
The quantity and kinetics of ice formation associated with cooling
of H. crispae from Vellerup Vig (20ppt) in active and P1 stages
were studied by differential scanning calorimetry (DSC). Groups
of 40–75 animals in each respective stage were transferred to 30μl
aluminum DSC pans. In order to avoid dehydrating the animals
during the removal of external water, the tardigrades were clumped
in the central part of the DSC pan and excess water was subsequently
removed with small pieces of delicate task wipes. Sample mass was
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Cyclomorphosis in tardigrades
2805
determined gravimetrically to the nearest 0.01mg using a fine-scale
AT261 Deltarange (Mettler-Toledo, Columbus, OH, USA), yielding
a total mass of 0.16–0.39mg (wet mass). The pans were sealed and
transferred to a calorimeter (Perkin Elmer DSC 7 equipped with an
Intercooler II mechanical cooling device), with an empty pan as
reference. The calorimeter was calibrated with gallium [melting
point, T m =29.78°C; melting enthalpy, H m =80.1 J g –1 ], water
(T m =0°C) and n-decane (T m =–29.66°C). All scans involved cooling
from 5°C to –40°C and subsequent reheating to 5°C at a cooling
rate of 5°Cmin –1 . Samples were reweighed following the DSC run
to ensure that no water loss had occurred. In order to determine the
water content of samples following the freeze/thaw cycle, pans were
punctured and dried at 80°C to a constant mass (dry mass). A
minimum of three groups of animals in each stage was used (see
Table1). The obtained thermograms (heat flows vs temperature)
were analyzed with respect to crystallization temperature (T c ),
amount of ice formed during the freezing exotherm (assuming that
the latent heat of crystallization is the same as for pure water), T m
and the osmotic pressure of the extracellular fluids as calculated by
the standard DSC 7 software. Ice contents were calculated using
the water content and the enthalpy of the freeze exotherm. The
temperature dependence of the enthalpy of crystallization of water
was taken into account as previously described (Kristiansen and
Westh, 1991). For hemolymph osmolality calculations, the onset
melting point measured by the DSC 7 software was determined
according to the approach of Nicholajsen and Hvidt (Nicholajsen
and Hvidt, 1994), in which the established melting point of the body
fluid was derived from a standard curve made from predetermined
NaCl solutions.
Preparation of experimental solutions
Salt water solutions of different osmotic pressure were made by
successive dilution with distilled water or by evaporative reduction
of 100% SW from the locality. Measurements of osmotic pressure
were made in parallel on a Vapro 5520 vapor pressure osmometer
(Wescor, Logan, UT, USA) and on a refractometer (S-1 Shibuya
Land, Tokyo, Japan).
Osmotic stress tolerance
Animals collected from the Danish as well as the Greenlandic
population were used for the current experiment. Groups of 20
specimens were transferred to small glass vials containing 4ml of
100% SW at 4°C, and specimens were either exposed to a gradual
increase or decrease in salinity. The gradual changes in salinity were
performed over the course of 4–5h by periodically replacing small
volumes of SW with prefixed solutions of either a higher or lower
salinity. Animal activity was concomitantly assessed. Animals were
allowed a period of 20–40min of acclimatization following a salinity
change prior to assessment. Individuals responsive to tactile stimuli
were considered active. Five groups of specimens in active and P1
stages were assessed at both hypo- and hyperosmotic salinities.
Volume measurements
Individual adult active-stage specimens of H. crispae (size
300–500μm) from Vellerup Vig (20ppt) were visualized in an
Olympus BX 51 microscope (Olympus), photographed using a
digital camera (C-5050, Olympus) and subsequently exposed for
set time periods of 30min, 1, 2, 4, 24 and 48h to saltwater solutions
with salinities of 2ppt, 10ppt and 40ppt. The osmotic treatments
were conducted in small glass vials containing 4ml of SW at 4°C.
At the end of each time interval, individuals were transferred, in a
drop of the appropriate solution, to glass microscope slides and
photographed under cover slips for subsequent estimations of body
volume. During photography, great care was taken to minimize the
time spent by the animals under the cover slips, in order to avoid
evaporative water loss, which would alter the osmotic pressure of
the solution. The animals were ensured total freedom of movement.
Following photography, individuals were returned to the respective
salinities until the end of the next set time period, when the process
was repeated. At each of the time intervals, 10–14 individuals were
photographed at each of the SW treatments. Images were analyzed
using DP-soft TM (Olympus), and total body volume was calculated
according to the equation: V total =π(r 2 bodyh body +2r 2 legh leg ), where V
is the volume of the specimen, r is the measured radius, and h is
the measured length of the body and hind legs, respectively.
In order to assess the behavioral response of H. crispae during
osmotic shock and to quantify potential mortality related to the
respective treatments, a separate experiment was performed.
Specimens (N=10) were transferred directly to 2ppt, 10ppt, 20ppt
(control) and 40 ppt, respectively, and animal activity was
subsequently monitored over the course of 48h at 4°C. Individuals
responsive to tactile stimuli at the above-mentioned set time periods
were considered active and alive (see Fig.7). Three groups exposed
to each treatment were assessed.
Measurement of hemolymph osmolality
Hemolymph osmolality was measured in individual tardigrades
following exposure for 30min, 4 and 48h to the experimental
solutions of 2ppt (62mOsmkg –1 ), 10ppt (311mOsmkg –1 ) and 40ppt
(1245 mOsm kg –1 ). Six animals were used for osmolality
determination in each of the experimental solutions. Six animals
kept at 20ppt (623mOsmkg –1 ) served as a control. Hemolymph
Table 1. Post-freeze survival and data obtained from differential scanning calorimetry on H. crispae from Vellerup Vig, Denmark (20 ppt)
in the active and P1 stages
Sample
Post-freeze
survival (%)
(frozen for 24 h)
Post-freeze
survival (%)
(frozen for
36 days)
Crystallization
temp. (°C)
Melting temp.
(°C)
Water content
(%)
Body-water
frozen during
freezing
exotherm (%)
Osmolality of
extracellular
–1
fluids (mOsm kg )
Pseudosimplex 1 53.3±15 (6) 12.7±7 (6) –19.6±3.1 (6) –4.29±0.79 (5) 67±4 (5) 59±3 (4) 928±77 (5)
Active 0 (6) 0 (6) –21.6±2.1 (3) –4.48±0.15 (3) 68±4 (3) 69±5 (3) 975±36 (3)
t-test (P0.05 (NS, not significant), P

2806
K. A. Halberg and others
samples (2–3nl) were collected by piercing individual specimens
under immersion oil (type A; 150 centistoke; Cargille Laboratories,
Cedar Grove, NJ, USA) using hand-pulled glass capillary tubes
(capacity 1 μl; Micro-caps, Drummond Scientific Company,
Broomall, PA, USA). Hemolymph samples were acquired through
capillary action and subsequently ejected into immersion oil. Care
was taken to ensure that the measurements were made on fluid
originating from hemolymph alone and samples containing gut
contents were discarded. Prior to sample collection, immersion oil
was collected into the capillary tube in order to avoid any evaporative
water loss. Using an Irvin loop, samples were immediately
transferred in a drop of immersion oil into sample oil wells (type
B; 1250 centistoke; Cargille Laboratories) of a calibrated nanoliter
osmometer (Clifton Technical Physics, Hartford, NY, USA), and
the osmolality (mOsmkg –1 ) was determined by melting point
depression (MDP=1.858°COsm –1 ).
Statistics
Significant differences between experimental and control conditions
were tested using unpaired, two-tailed t-tests, and a significance level
of P≤0.05.
RESULTS
Survival at sub-zero temperatures
The external morphology of H. crispae in the active and the P1
stage is shown in Fig.1. Notably, the P1 stage is characterized by
a conspicuous double cuticle, in which both the mouth and cloaca
are closed by cuticular thickenings (Fig.1A–C). Post-freeze survival
following both short- and long-term exposure to subzero
temperatures is listed in Table1. An example of the quantitative
kinetics of ice formation associated with the freeze/thaw cycle of
P1-stage H. crispae kept at 20ppt is illustrated in Fig.2. The
collective DSC analysis of the onsets and areas of peaks after the
freeze/thaw cycles, together with the water content of the samples,
provide the remaining results listed in Table1.
Whereas animals in the active stage and P2 stage (data not shown)
were intolerant of freezing, animals in the P1 stage were
demonstrated to be freeze-tolerant. Post-freeze survival following
24h exposure to –20°C was 53.3±15%; however, when prolonging
the period spent at subzero temperatures to 36days, the survival of
animals in the P1 stage decreased to 12.7±7% (Table1). Animal
recovery was monitored over a period of 96 h following the
freeze/thaw cycle; however, the majority of animals had resumed
activity after a period of 48h. No additional recovery was monitored
beyond 96h following any of the investigated treatments.
Substantial ice formation proceeded rapidly following the first
ice nucleation in the freeze-tolerant P1 stage with a mean of
–19.6±3.1°C; as indicated by the large exotherm in Fig.2. At the
given salinity, the amount of ice formed during the freezing
exotherm amounted to approximately 60% of the body water
(Table1). The absence of additional small exothermic peaks during
the subsequent cooling to –40°C indicated that no additional ice
formation occurred following the initial large freezing exotherm.
The freeze exotherm lasted less than one minute. The initial
separation of ice (T c ) occurred in the temperature range of –15.4 to
–23.2°C.
Only marginal differences in T c of animals in the P1 and active
stage were observed (Table1), suggesting an absence of seasonal
variations in ice-nucleating activity in H. crispae. In spite of invariant
water contents between the two stages, the amount of water
crystallizing during cooling in animals in the active stage was
significantly higher than in animals in the P1 stage; however, the
melting points of the two stages remained largely unaltered. The
latter indicates that body fluid osmolality at a given external salinity
is unaffected by the animal’s transition from the active to the P1
stage.
Volume- and osmoregulatory capacity
Fig.3 shows the percentage of active animals of H. crispae following
the exposure to gradual changes in the external salinity. When
comparing Danish P1 and active-stage H. crispae, the active stage
displayed a larger tolerance towards the more concentrated SW
solutions and were slightly more tolerant of the very dilute solutions.
Indeed, a significantly higher percentage of active-stage specimens
Differential heat flow (mW)
15
7.5
Exotherm
Cooling
Heating
Endotherm
0
–40 –35 –30 –25 –20 –15 –10 –5 0 5
Temperature (°C)
Fig. 2. Representative thermogram displaying the exothermic and
endothermic events associated with the cooling and heating of a sample
containing P1-stage Halobiotus crispae from Vellerup Vig, Denmark
(20 ppt). The crystallization and melting temperatures were estimated as
the onsets of peaks (–17.5°C and –4.5°C, respectively) using the DSC 7
software (see also Table 1). The very low crystallization temperatures
measured exclude the presence of physiologically relevant ice-nucleating
agents.
Activity (%)
100
80
60
40
20
0
Active, Vellerup
P1, Vellerup
Active, Nipisat
Osmolality (mOsm kg –1 )
0 500 1000 1500 2000 2500
*
*
0 10 20 30 40 50 60 70 80 90
Salinity (ppt)
100
Fig. 3. Osmotic stress tolerance of Halobiotus crispae in the P1 and active
stages. Active-stage () and P1-stage () specimens from the Danish
population at Vellerup Vig. Active-stage () specimens from the
Greenlandic population at Nipisat Bay. Data are means ± s.e.m. from five
independent experiments. *, significantly different from Vellerup Vig P1
stage (P

Cyclomorphosis in tardigrades
2807
remained active in the salinity spectrum of 0–3ppt and 45–60ppt,
as compared with the animals in the P1 stage, and in general seemed
less affected by the impositions of osmotic stress. Our preliminary
data on P2 from Vellerup Vig show that this stage tolerates very
dilute solutions better than the other stages, yet is the least tolerant
of increases in salinity, becoming inactive at around 50ppt. As a
comparison, the active stage from Greenland (Nipisat Bay), living
in a more exposed habitat compared with the Vellerup Vig
population, displayed an even higher tolerance to concentrated SW,
with observed activity at 80ppt.
In the following, we investigate in detail volume and
osmoregulation in active-stage H. crispae from Vellerup Vig kept
at a control salinity of 20ppt (Figs4–8). When exposed to a severe
hypo-osmotic shock of 2 ppt (63 mOsm kg –1 ), the animals
exhibited a large significant increase in body volume, resulting
in a total body volume of 127±11% after merely 0.5h of exposure
(Fig.4A,B). During this time period, animals became bloated and
rigid and most specimens lost locomotory functions (Fig.7A).
This passive uptake of water continued during the initial 2h of
the exposure, culminating in a total body volume of 162±17%.
However, after this period of time, a regulatory volume decrease
(RVD) was observed. The total body volume of specimens was
considerably reduced to 114±14% following 48h immersion and
was not significantly different from the controls (Fig. 4B).
Additionally, an increase in the number of active animals was
observed, yet a considerable number of specimens remained
passive throughout the treatment (Fig. 7A). Nevertheless,
following a gradual return to 20 ppt, all animals regained
locomotory functions. No mortality was observed in any of the
treatments.
Upon immersion of individual specimens into a less severe hypoosmotic
media of 10ppt (311mOsmkg –1 ) a similarly significant
increase in total body volume was observed; reaching a mean value
of 132±11% after 0.5h exposure (Fig.5A,B). However, following
this initial increase, total body volume stabilized, and a RVD was
observed after 1–2h exposure. After 4h incubation, total body
volume was 110±8%, which was not significantly different from
the control situation. An effect on the locomotory functions was
observed initially, as some animals displayed sluggish movements,
yet only a limited number of animals were passive during this
experiment (Fig.7B).
When transferring H. crispae to a hyperosmotic solution of 40ppt
(1245mOsmkg –1 ), a significant decrease in total body volume was
observed (Fig.6A,B). Total body volume was significantly reduced
to 66±9% following the first 0.5h of immersion and remained largely
unaltered during the following hours of the treatment. After 24h,
most specimens had displayed a regulatory volume increase,
resulting in a mean total body volume of 82±9%, yet total body
volume remained significantly different from the control situation
even after 48h. Nevertheless, animal motility was little affected by
the hypertonic shock, neither at the initial transfer nor throughout
the rest of the experiment (Fig.7C).
20 ppt,
2 ppt,
623 mOsm kg –1 62 mOsm kg –1
A
0h 0.5h 1h 2h 4h 24h 48h
20 ppt,
10 ppt,
623 mOsm kg –1 311 mOsm kg –1
A
0h 0.5h 1h 2h 4h 24h 48h
B
Total body volume (%)
200
180
160
(12)
(6)
140
(12)
120
100
80
60
40
(6)
(12)
(12)
(6)
(10)
0 1 2 3 4 20 30 40 50
Exposure time (h)
(10)
(6)
1400
1200
1000
800
600
400
200
Hemolymph osmotic pressure (mOsm kg –1 )
B
Total body volume (%)
200
180
160
140
120
100
80
60
40
(6)(10)(12) (12)
(10) (13)
(6)
(6)
(11)
0 1 2 3 4 20 30 40 50
Exposure time (h)
(6)
1400
1200
1000
800
600
400
200
Hemolymph osmotic pressure (mOsm kg –1 )
Fig. 4. (A) Halobiotus crispae in active stage from Vellerup Vig, Denmark
(20 ppt). Light-microscopical images at different time points following
exposure to 2 ppt (62 mOsm kg –1 ) of a single specimen (scale bar=100 μm).
(B) Changes in total body volume () and measured internal osmolality ()
over a period of 48 h following exposure to an external salinity of 2 ppt.
Data are expressed as means ± s.d. Numbers in parentheses indicate the
number of animals used for assessment of body volume and hemolymph
osmolality at each time point.
Fig. 5. (A) Halobiotus crispae in active stage from Vellerup Vig, Denmark
(20 ppt). Light-microscopical images at different time points following
exposure to 10 ppt (311 mOsm kg –1 ) of a single specimen (scale
bar=100 μm). (B) Changes in total body volume () and measured internal
osmolality () over a period of 48 h following an exposure to an external
salinity of 10 ppt. Data are expressed as means ± s.d. Numbers in
parentheses indicate the number of animals used for assessment of body
volume and hemolymph osmolality at each time point.
THE JOURNAL OF EXPERIMENTAL BIOLOGY

2808
K. A. Halberg and others
20 ppt,
40 ppt,
623 mOsm kg –1 1245 mOsm kg –1
A
B
Total body volume (%)
0h 0.5h 1h 2h 4h 24h 48h
200
180
160
140
120
100
80
60
(6)
(6)
(12)
(12) (14)
(11)
(13) (11)
1400
1200
1000
200
40
0 1 2 3 4 20 30 40 50
Exposure time (h)
Fig. 6 (A) Halobiotus crispae in active stage from Vellerup Vig, Denmark
(20 ppt). Light-microscopical images at different time points following
exposure to 40 ppt (1245 mOsm kg –1 ) of a single specimen (scale
bar=100 μm). (B) Changes in total body volume () and measured internal
osmolality () over a period of 48 h following an exposure to an external
salinity of 40 ppt. Data are expressed as means ± s.d. Numbers in
parentheses indicate the number of animals used for assessment of body
volume and hemolymph osmolality at each time point.
Notably, hemolymph osmotic pressure differed significantly
from the control condition at all time points examined during the
various salinity treatments (Figs4–6). Hemolymph osmolality of H.
crispae varied in proportion to the gradient set up by the salinity
transfer. The body fluids of animals kept under control conditions
(20ppt, 623mOsmkg –1 ) had an osmolality of 926±29mOsmkg –1 ,
(6)
(6)
800
600
400
Hemolymph osmotic pressure (mOsm kg –1 )
indicating that H. crispae hyper-regulates during steady-state
conditions. This hyperosmotic regulation was independently
confirmed by the DSC investigation in which the hemolymph
osmolality was measured at 975±36mOsmkg –1 (Table1). The two
measurements are not significantly different (t-test, P

Cyclomorphosis in tardigrades
2809
Hemolymph osmolality (mOsm kg –1 )
1400
1200
1000
800
600
400
200
Active stage
Osmotic performance
Isoosmotic line
0
0 200 400 600 800 1000 1200 1400
External osmolality (mOsm kg –1 )
Fig. 8. Measured hemolymph osmolality of Halobiotus crispae (active stage
from Vellerup Vig, Denmark) during steady-state conditions after 48 h
acclimation to 2 ppt (63 mOsm kg –1 ), 10 ppt (311 mOsm kg –1 ), 20 ppt
(623 mOsm kg –1 ) and 40 ppt (1245 mOsm kg –1 ), respectively. Each point
represents the mean ± s.d. of the individual experiments. The broken line
indicates the isoosmotic line at which no osmoregulation occurs.
functionally characterized as a movable cyst (Kristensen, 1982;
Møbjerg et al., 2007; Guidetti et al., 2008). In Denmark (Vellerup
Vig), this stage is dominant during the summer months, presumably
enabling H. crispae to withstand heat stress and oxygen depletion.
The active stage, the only stage at which active feeding and sexual
reproduction occur, is the dominant stage during the Greenlandic
summer, whereas this stage is present during late winter and the
spring months in Denmark (Møbjerg et al., 2007).
Freeze avoidance and freeze tolerance
Winter temperatures are frequently below the equilibrium freezing
point of the surrounding seawater at least in some portions of the
natural environments of H. crispae, and certain habitats may even
become completely frozen for extended periods of the year
(Kristensen, 1982). Enduring such hostile surroundings requires
corresponding cold-tolerance strategies that enable long-term
survival. Traditionally, two main options are exploited by
ectothermic animals when faced with subzero temperatures, i.e.
freeze avoidance and freeze tolerance (Lee, 1991). When exposing
animals in the P1 stage of H. crispae to temperatures below the
equilibrium freezing point (T c ) of their body fluids, freeze tolerance
is demonstrated, indicating that winter survival could involve
extracellular ice formation in this species. However, the finding that
mortality increases with prolonged exposure to subzero temperatures
suggests that the consequent damages accumulate in proportion to
the time spent exposed to freezing conditions. This observation is
likely explained by the depletion of essential metabolites and is of
particular interest in Arctic habitats in which subfreezing
temperatures have to be endured for long periods of time.
In freeze-tolerant organisms, ice formation is usually promoted
at relatively high subzero temperatures (–2 to –10°C) by icenucleating
agents present in the extracellular fluid (Zachariassen,
1985; Block, 1991; Westh and Kristensen, 1992). The adaptive
advantage of such a strategy is that the process of ice formation
proceeds relatively slowly at relatively high temperatures, enabling
the organism to maintain the damage associated with freezing within
tolerable boundaries. Indeed, both the localization and the amount
of ice formed in freeze-tolerant organisms are usually under tight
control. Surprisingly, our calorimetric investigation of the freezetolerant
P1 stage reveals that ice crystallization occurs at
approximately –20°C, excluding the presence of any physiologically
relevant ice-nucleating agents in this stage. In fact, the very low
crystallization temperatures measured in both stages suggest that
the capacity for supercooling is maintained throughout the majority
of the year, which is in general contrast to the pattern observed in
most other freeze-tolerant invertebrates (Block, 1991; Westh and
Kristensen, 1992; Ramløv et al., 1996). However, in spite of an
invariant melting point and water content between the two stages,
the amount of water crystallized during cooling to –40°C appears
to be about 60% for animals in the P1 stage and 70% for animals
in the active stage (see Table 1). Consequently, the cellular
dehydration induced by freezing is expected to be much higher in
animals in the active stage, compared with that of specimens in the
P1 stage. This reduction in ice accumulation in specimens in the
P1 stage could potentially be explained by an increased production
of macromolecules [which kinetically inhibits ice formation but has
negligible effect on the melting temperature (see Westh and
Kristensen, 1992)], as compared with the active stage. Whether this
observation alone explains the observed freeze tolerance is difficult
to determine. Selected cryptobiotic species of tardigrades, nematodes
as well as some freeze-tolerant insects tolerate as much as 80% of
the body water being converted into ice (Westh and Kristensen,
1992; Ramløv and Westh, 1993; Wharton and Block, 1997;
Hengherr et al., 2009).
The apparent morphological difference between the two stages
is similarly relevant in regard to the observed freeze tolerance. The
P1 stage is formed from an incomplete molt in which both the mouth
and cloaca become sealed by cuticular thickenings (see Fig.1), and
the gut content is often shed prior to this transition. In nature, ice
nucleation can be initiated by a wide range of exogenous substances
(Wharton and Worland, 1998). Consequently, the additional layer
of cuticle could increase the capacity to avoid inoculative freezing
in animals in the P1 stage, as has been demonstrated for eggs of
the nematode Panagrolaimus davidi Timm 1971 (see Wharton,
1994). Indeed, the extensive capacity for supercooling in both the
active and P1 stage, along with the additional layer of cuticle and
the clearing of gut contents in P1, would indicate that H. crispae
preferentially seek to avoid internal ice formation. Nevertheless,
animals in the P1 stage tolerate internal ice formation for both shorter
and longer periods of time.
Volume and osmoregulation
Our results indicate that active-stage H. crispae is the most tolerant
of changes in external salinity. Specimens from the population from
Nipisat Bay, Greenland exhibit an increased tolerance towards
concentrated SW solutions as compared with animals from Vellerup
Vig, Denmark, suggesting that the ability to tolerate large increases
in salinity is potentiated by living in a more-exposed habitat. The
observed volume regulatory response of active-stage H. crispae
during hypo- and hypertonic treatments differs in a significant way.
When exposed to the hypo-osmotic solutions, the initial increase in
total body volume was regulated to a new steady state, which was
not significantly different from the control condition. In fact, a new
steady-state value was demonstrated after merely 4h immersion in
the external medium of 10ppt. Conversely, during acute exposure
to concentrated seawater (40ppt), a partial recovery to normal levels
was demonstrated; however, total body volume remained
significantly different from the control condition even after 48h
immersion. These data suggest that the body volume of H. crispae
is more tightly regulated during exposure to dilute as compared with
THE JOURNAL OF EXPERIMENTAL BIOLOGY

2810
K. A. Halberg and others
more concentrated saltwater solutions. Interestingly, this observation
seems reflected in an evolutionary context. According to our
previous study, Halobiotus has evolved within the freshwater genus
Isohypsibius, thus potentially explaining the enhanced volume
regulatory response during exposure to dilute media (Møbjerg et
al., 2007).
When submitted to osmotic shock of 10 ppt and 40 ppt,
respectively, our data show that H. crispae experience few
limitations in terms of motility. Active-stage specimens from the
Nipisat population even retain activity when exposed to a gradual
salinity increase to 60ppt (Fig.3). However, upon direct transfer to
an extreme seawater dilution of 2ppt, animal activity is markedly
reduced, probably due to the pronounced increase in hydrostatic
pressure and concomitant reduction in hemolymph osmotic pressure.
Proper locomotory function in tardigrades relies on the hydrostatic
pressure of the body cavity (Kinchin, 1994); thus, maintaining an
appropriate body volume is essential to normal coordination of
movement. In addition, as the membrane potentials of animal cells
are highly dependent upon extracellular ionic strength (Spyropoulos
and Teorell, 1968), the concomitant changes in hemolymph osmotic
pressure could influence animal motility due to inhibition of neuromuscular
activity. Indeed, the fact that a significant number of
animals were observed passive during exposure to 2ppt, while
animals remained largely unaffected during exposure to 10ppt, in
spite of experiencing comparable average changes in body volume
(compare Fig.4B and Fig.5B after 0.5h immersion), suggests that
not only total body volume but also hemolymph osmolality is an
important factor in maintaining locomotory functions.
Exposing H. crispae to severe osmotic stress reveals that this
species is a euryhaline osmoconformer, in which the hemolymph
osmotic pressure is largely governed by the external environment.
However, when analyzing hemolymph osmotic pressure at steady
state following 48h exposure to the various salinity treatments as
a function of the external osmolality, an interesting pattern emerges.
H. crispae maintains a large osmotic pressure gradient between the
internal and external environment, thus distinctly hyper-regulating
during all investigated salinity treatments – albeit markedly less in
concentrated seawater. This would imply a large water turnover in
this animal, with osmotic water uptake being balanced by the
excretion of dilute urine.
Hyperosmoregulation is known in other euryhaline invertebrates.
The crayfish Procambarus clarkii Girard 1852 (Arthropoda)
produces highly dilute urine and is a strong hyperosmoregulator in
freshwater (Sarver et al., 1994). However, the excreted urine
becomes progressively more concentrated in media of higher ionic
strength and is nearly isoosmotic when exposed to an external
concentration of 750mOsmkg –1 , at which P. clarkii cease to hyperregulate
(Sarver et al., 1994). Moreover, similar osmoregulatory
responses have been reported from nematodes (Fusé et al., 1993;
Forster, 1998). Indeed, the internal osmolality of the parasitic
nematode Pseudoterranova decipiens (Krabbe, 1878) was
maintained 90mOsmkg –1 above that of the external environment
during exposure to media of widely varying osmolality (Fusé et al.,
1993).
Three glands positioned at the transition zone between the
midgut and rectum of eutardigrades are traditionally ascribed an
osmoregulatory function. The term used for these structures, i.e.
Malpighian tubules, was introduced more than a century ago (Plate,
1889). The positional conformity of the Malpighian tubules in
eutardigrades and in hexapods has been used as a strong argument
in favor of a homology between these structures (Greven, 1982;
Møbjerg and Dahl, 1996). However, at present, no functional data
exist relating the Malpighian tubules of tardigrades to an
osmoregulatory role. Nevertheless, several detailed morphological
investigations of the tubules support the hypothesis. These studies
have provided ultrastructural data, which are in agreement with an
active transporting epithelium involved in solute and fluid transport
(Greven, 1979; Weglarska, 1987a; Weglarska, 1987b; Møbjerg and
Dahl, 1996; Peltzer et al., 2007). As holds for insects, the Malpighian
tubules of tardigrades are considered secretion–reabsorption kidneys.
In light of the ultrastructural data available on tardigrade Malpighian
tubules, it seems reasonable to assume that the first steps in urine
formation take place across initial segment cells, characterized by
a conspicuous basal labyrinth, numerous mitochondria and an
enlarged apical surface. There is, moreover, ultrastructural support
for assigning the tardigrade rectum an osmoregulatory function
(Dewel and Dewel, 1979). A possible mode of urine formation was
outlined by Dewel and Dewel (Dewel and Dewel, 1979). They
suggest that isoosmotic urine produced by the Malpighian tubules
is modified in the rectum through the active reabsorption of solutes,
leading to the excretion of hypo-osmotic urine. Our data seem in
favor of such a mechanism of urine formation. Interestingly,
preliminary and unpublished data (H.R. research group) on
Richtersius coronifer (Richters 1903) indicate that this species also
remains hyperosmotic during exposures to a range of external
salinities and it is therefore likely that hyper-regulation, and possibly
hypo-osmotic urine formation, is a general feature of eutardigrades.
However, until functional studies at the cellular and molecular level
are performed, the exact mechanisms involved in osmoregulation
in tardigrades remain to be elucidated.
In conclusion, we show that the transition between the individual
cyclomorphic stages of H. crispae is associated with profound
changes in the physiology of the animal. Our results show that
animals in the active stage tolerate large changes in the external
osmotic pressure by regulating their total body volume and by
enduring large concomitant changes in hemolymph osmotic
pressure. H. crispae remains hyperosmotic at any investigated
external salinity, suggesting that this species is a strong hyperregulator.
Our study is the first to provide evidence for the volume
and osmoregulatory capacity in Tardigrada. Whereas animals in the
active stage are intolerant of freezing, the P1 stage is demonstrated
to be freeze tolerant. The relatively low crystallization temperature
reveals that extensive supercooling of the body fluids takes place
during cooling and that no physiologically relevant ice-nucleating
agents are present.
We would like to thank the crew onboard the research vessel R/W Porsild (Arctic
Station, Qeqertarsuaq, Greenland) who made the collection at the type locality,
Nipisat, possible. Funding came from the Carlsberg Foundation and from the
2008 Faculty of Science, University of Copenhagen Freja-Programme.
DSC
MDP
RVD
SW
T c
T m
LIST OF ABBREVIATIONS
differential scanning calorimetry
melting point depression
regulatory volume decrease
seawater
crystallization temperature
melting temperature
REFERENCES
Block, W. (1991). To freeze or not to freeze Invertebrate survival of sub-zero
temperatures. Funct. Ecol. 5, 284-290.
Convey, P. and McInnes, S. J. (2005). Exceptional tardigrade-dominated ecosystems
in Ellsworth Land, Antarctica. Ecology 86, 519-527.
Dewel, R. A. and Dewel, W. C. (1979). Studies on the tardigrades. J. Morphol. 161,
79-110.
Forster, S. J. (1998). Osmotic stress tolerance and osmoregulation of intertidal and
subtidal nematodes. J. Exp. Mar. Biol. Ecol. 224, 109-125.
THE JOURNAL OF EXPERIMENTAL BIOLOGY

Cyclomorphosis in tardigrades
2811
Fusé, M., Davey, K. G. and Sommerville, R. I. (1993). Osmoregulation in the
parasitic nematode Pseudoterranova decipiens. J. Exp. Biol. 175, 127-142.
Garey, J. R., Krotec, M., Nelson, D. R. and Brooks, J. (1996). Molecular analysis
supports a tardigrade-arthropod association. Invertebr. Biol. 115, 79-88.
Giribet, G., Carranza, S., Baguña, J., Riutort, M. and Ribera, C. (1996). First
molecular evidence of the existence of a Tardigrada+Arthropoda clade. Mol. Biol.
Evol. 13, 76-84.
Greven, H. (1979). Notes on the structure of vasa Malpihgii in the eutardigrade
Isohypsibius augusti (Murray, 1907) Zesz. Nauk. Uniw. Jagiel. Prace Zool. 25, 87-
95.
Greven, H. (1982). Homologues or analogues A survey of some structural patterns in
Tardigrada. In Proceedings of the Third International Symposium on the Tardigrada
(ed. D. R. Nelson), pp. 55-76. Johnson City, TN: East Tennessee State University
Press.
Grøngaard, A., Pugh, P. J. A. and McInnes, S. (1999). Tardigrades, and other
cryoconite biota, on the Greenland ice sheet. Zool. Anz. 238, 211-214.
Guidetti, R., Boschini, D., Altiero, T., Bertolani, R. and Rebecchi, L. (2008).
Diapause in tardigrades: a study of factors involved in encystment. J. Exp. Biol. 211,
2296-2302.
Hengherr, S., Heyer, A. G., Köhler, H. R. and Schill, R. O. (2008). Trehalose and
anhydrobiosis in tardigrades: evidence for divergence in response to dehydration.
FEBS J. 275, 281-288.
Hengherr, S., Worland, M. R., Reuner, A., Brümmer, F. and Schill, R. O. (2009).
Freeze tolerance, supercooling pionts and ice formation: comparative studies on the
subzero temperature survival of limno-terrestrial tardigrades. J. Exp. Biol. 212, 802-
807.
Horikawa, D. D., Sakashita, T., Katagiri, C., Watanabe, M., Kikawada, T.,
Nakahara, Y., Hamada, N., Wada, S., Funayama, T., Higishi, S. et al. (2006).
Radiation tolerance in the tardigrade Milnesium tardigradum. Int. J. Radiat. Biol. 82,
843-848.
Jönson, K. I. and Schill, R. O. (2007). Induction of Hsp70 by dessication, ionizing
radiation and heat-shock in the eutardigrade Richtersius coronifer. Comp. Biochem.
Physiol. 146B, 456-460.
Kinchin, I. M. (1994). The Biology of Tardigrades, pp. 1-186. London: Portland Press.
Kristensen, R. M. (1982). The first record of cyclomorphosis in Tardigrada based on a
new genus and species from Arctic meiobenthos. Z. Zool. Syst. Evol. Forsch. 20,
249-270.
Kristiansen, J. and Westh, P. (1991). Freezing behaviour of multilamellar vesicles in
0.9% sodium chloride. Cryo Letters 12, 167-176.
Lee, R. E. (1991). Principles of insect low temperature tolerance. In Insects at Low
Temperatures (ed. R. E. Lee and D. L. Denlinger), pp. 17-46. New York: Chapman
and Hall.
Mallatt, J. M., Garey, J. R. and Shultz, J. W. (2004). Ecdysozoan phylogeny and
Baysian inference: first use of nearly complete 28S and 18S rRNA gene sequences
to classify the arthropods and their kin. Mol. Phylogenet. Evol. 31, 178-191.
Møbjerg, N. and Dahl, C. (1996). Studies on the morphology and ultrastructure of the
Malpighian tubules of Halobiotus crispae Kristensen, 1982 (Eutardigrada). Zool. J.
Linn. Soc. 116, 85-99.
Møbjerg, N., Jørgensen, A., Eibye-Jacobsen, J., Halberg, K. A., Persson, D. and
Kristensen, R. M. (2007). New records on cyclomorphosis in the marine
eutardigrade Halobiotus crispae (Eutardigrada: Hypsibiidae). J. Limnol. 66 Suppl. 1,
132-140.
Nicholajsen, H. and Hvidt, Å. (1994). Phase behaviour of the system trehalose-NaCl-
Water. Cryobiology 31, 199-205.
Pelzer, B., Dastych, H. and Greven, H. (2007). The osmoregulatory/excretory organs
of the glacier-dwelling eutardigrade Hypsibius klebelsbergi Mihelcic, 1959
(Tardigrada). Mitt. Hamb. Zool. Mus. Inst. 104, 61-72.
Plate, L. H. (1889). Beiträge zur Naturgeschichte der Tardigraden. Zool. Jb. Anat.
Ontog. 3, 487-550.
Ramløv, H. and Westh, P. (1992). Survival of the cryptobiotic tardigrade Adorybiotus
coronifer during cooling to –196°C: effect of cooling rate, trehalose level and short
term pre-acclimation. Cryobiology 29, 125-130.
Ramløv, H. and Westh, P. (1993). Ice formation in the freeze tolerant alpine weta
Hemideina maori Huttton (Orthoptera; Stenopelmatidae). Cryo Letters 14, 169-176.
Ramløv, H. and Westh, P. (2001). Cryptobiosis in the Eutardigrade Adorybiotus
(Richtersius) coronifer: tolerance to alcohols, temperature and de novo protein
synthesis. Zool. Anz. 240, 517-523.
Ramløv, H., Wharton, D. A. and Wilson, P. W. (1996). Recrystalization in a freezingtolerant
Antarctic nematode Panagrolaimus davidi, and an alpine weta Hemideina
maori (Orthoptera; Stenopelmatidae). Cryobiology 33, 607-613.
Sarver, R. G., Flynn, M. A. and Holliday, C. W. (1994). Renal Na, K-ATPase and
osmoregulation in the crayfish Procambarus clarkii. Comp. Biochem. Physiol. 107A,
349-356.
Schill, R. O., Steinbrück, G. H. B. and Köhler, H. R. (2004). Stress gene (hsp70)
sequences and quantitative expression in Milnesium tardigradum (Tardigrada) during
active and cryptobiotic stages. J. Exp. Biol. 207, 1607-1613.
Spyropoulos, C. S. and Teorell, T. (1968). The dependence of nerve membrane
potentials upon extracellular ionic strength. Proc. Natl. Acad. Sci. USA 60, 118-125.
Weglarska, B. (1987a). Studies on the excretory system of Isohypsibius granulifer
Thulin (Eutardigrada). In Biology of Tardigrades: Selected Symposia and
Monographs 1 (ed. R. Bertolani), pp. 15-24. Modena: U.Z.I. Mucchi.
Weglarska, B. (1987b). Morphology and ultrastructure of the excretory system in
Dactylobiotus dispar (Murray) (Eutardigrada). In Biology of Tardigrades: Selected
Symposia and Monographs 1 (ed. R. Bertolani), pp. 25-33. Modena: U.Z.I. Mucchi.
Westh, P. and Kristensen, R. M. (1992). Ice formation in the freeze-tolerant
eutardigrades Adorybiotus coronifer and Amphibolus nebulosus studied by
differential scanning calorimetry. Polar Biol. 12, 693-699.
Wharton, D. A. (1994). Freezing avoidance in the eggs of the Antarctic nematode
Panagrolaimus davidi. Fundam. Appl. Nematol. 17, 239-243.
Wharton, D. A. and Block, W. (1997). Differential scanning calorimetry studies on an
Antarctic nematode (Panagrolaimus davidi) which survives intracellular freezing.
Cryobiology 34, 114-121.
Wharton, D. A. and Worland, M. R. (1998). Ice nucleating activity in the freezingtolerant
Antarctic nematode Panagrolaimus davidi. Cryobiology 36, 279-286.
Zachariassen, K. E. (1985). Physiology of cold tolerance in insects. Physiol. Rev. 65,
799-831.
THE JOURNAL OF EXPERIMENTAL BIOLOGY

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