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Graduate School
2013
The Role of Polyamines in Osmotic Stress
Tolerance in Gulf Killifish Fundulus Grandis
Ying Guan
Louisiana State University and Agricultural and Mechanical College, [email protected]
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Recommended Citation
Guan, Ying, "The Role of Polyamines in Osmotic Stress Tolerance in Gulf Killifish Fundulus Grandis" (2013). LSU Doctoral
Dissertations. 1123.
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THE ROLE OF POLYAMINES IN OSMOTIC STRESS TOLERANCE IN GULF
KILLIFISH FUNDULUS GRANDIS
A Dissertation
Submitted to the Graduate Faculty of the
Louisiana State University and
Agricultural and Mechanical College
in partial fulfillment of the
requirements for the degree of
Doctor of Philosophy
in
The Department of Biological Sciences
by
Ying Guan
B.S., Qiqihaer Medical School, 2003
M.S., Jinan University, 2006
December 2013
ACKNOWLEDGMENTS
I would like to express my deep appreciation and gratitude to my advisor, Dr.
Fernando Galvez, for the patient guidance and mentorship he provided all the way from
the beginning to the completion of the Ph.D program. I am truly fortunate to have the
opportunity to work with him.
I would also like to thank my committee members; Dr. William B. Stickle, Dr.
Steve C. Hand, Dr. Joseph F. Siebenaller; for their friendly guidance, thought-provoking
suggestions, and for the general collegiality that each of them offered to me over the
years.
I would like to thank Dr. Benjamin Dubansky, Yanling Meng, Christine
Savolainen, Charles Brown, and my friends in the Department of Biological Sciences.
Their friendship and help with my experiments made my time in the lab much easier.
I give special thanks to my husband, Guangming Hu, for his innumerable
sacrifices in shouldering far more than his fair share of the parenting and financial
burdens.
Lastly, my sincere thanks go to my parents, Tang and Feng. They have been there
for me from day one. Thank you for all of the love, support, encouragement and
dedication.
ii
TABLE OF CONTENTS
ACKNOWLEDGMENTS .................................................................................................. ii
ABSTRACT ....................................................................................................................... iv
CHAPTER 1: LITERATURE REVIEW ............................................................................ 1
1.1 General principles of osmoregulation in teleost fish ................................................. 1
1.2 Euryhaline Fundulus species – model for environmental salinity tolerance ............ 2
1.3 The influence of environmental salinity on fish gill remodeling .............................. 2
1.4 Transcriptional and enzymatic activities of key enzymes ......................................... 6
1.5 Physiological function of polyamines to salinity challenge .................................... 13
1.6 Role of polyamines in cell proliferation and cell apoptosis .................................... 14
1.7 Brief outline of main points of each chapter ........................................................... 15
CHAPTER 2: THE INFLUENCE OF HYPOOSMOTIC EXPOSURE ON
POLYAMINE BIOSYNTHESIS IN THE GULF KILLIFISH ........................................ 18
2.1 Introduction ............................................................................................................. 18
2.2 Materials and methods ............................................................................................ 20
2.3 Results ..................................................................................................................... 27
2.4 Discussion ............................................................................................................... 37
CHAPTER 3: THE POTENTIAL ROLE OF POLYAMINES IN GILL
EPITHELIAL REMODELING DURING EXTREME HYPOOSMOTIC
CHALLENGES IN THE GULF KILLIFISH ................................................................... 43
3.1 Introduction ............................................................................................................. 43
3.2 Materials and methods ............................................................................................ 46
3.3 Results ..................................................................................................................... 54
3.4 Discussion ............................................................................................................... 64
CHAPTER 4: THE INFLUENCE OF OSMOTIC STRESS ON
TRANSCRIPTION AND TRANSLATION OF ARGINASE, ORNITHINE
DECARBOXYLASE, AND POLYAMINE OXIDASE IN TISSUES ............................ 72
4.1 Introduction ............................................................................................................. 72
4.2 Materials and methods ............................................................................................ 73
4.3 Results ..................................................................................................................... 77
4.4. Discussion .............................................................................................................. 83
CHAPTER 5: CONCLUSION ......................................................................................... 88
REFERENCES ................................................................................................................. 94
APPENDIX: PERMISSION OF REPRINT ................................................................... 111
VITA ............................................................................................................................... 112
iii
ABSTRACT
The main objective of this study was to describe the mRNA levels and enzymatic
activities of key enzymes for polyamine metabolism, to measure polyamine levels, and to
assess putative roles of polyamines in the gills of the Fundulus grandis during
hypoosmotic challenge. The influence of irreversible inhibition of Odc by alpha-DLdifluoromethylornithine (DFMO) was also assessed in the gills. The mRNA levels and
enzymatic activities of Arg, Odc and Pao was assessed in other tissues during
hypoosmoitc challenges.
Adult F. grandis were reared in 5 ppt and acutely transferred to 5, 2, 1, 0.5, and
0.1 ppt water, and gills were sampled at 6 h, 1 d, 3 d, and 7 d post-transfer. Hypoosmotic
exposure produced increases in the arg II and odc mRNA levels, in gill Odc activity, and
in the concentrations of putrescine, spermidine, and spermine. DFMO application
inhibited Odc activity and reduced polyamine levels after0.1 ppt transfer. The ratio of
putrescine level over the sum levels of spermidine and spermine increased after 0.1 ppt
exposure at 1 d and beyond. Concomitant with freshwater acclimation, an increase in
Pao activity suggested that polyamine catabolism was upregulated in the gills. The
phenotype of mitochondrion-rich cells (MRCs) in the gill epithelium shifted from a
seawater type to a freshwater type following transfer to 0.1 ppt water in correlation with
the increase in mRNA levels of arg II and odc in MRCs. In addition, the isolated
opercular epithelium pretreated with spermidine had a lower active Cl- secretion rate and
membrane conductance following symmetrical hypotonic exposure. The mRNA levels
and enzymatic activities of Arg II, Odc, and Pao were upregulated in the intestine and
liver during hypoosmotic exposure, suggesting that polyamine levels are regulated in
iv
multiple tissues of the killifish. The putative roles of polyamines include inducing cell
apoptosis by increasing caspase-3 activity, stimulating cell proliferation by increasing the
levels of c-fos and c-myc mRNA levels, and inducing cell swelling via the modulation of
Cl- secretion in the gills following hypoosmotic challenges.
In summary, highly cationic polyamines mediate early phase compensatory
responses in the euryhaline killifish when faced with osmotic challenges.
v
CHAPTER 1: LITERATURE REVIEW
1.1 General principles of osmoregulation in teleost fish
Teleost fish regulate the ionic strength and composition of their extracellular
fluids (ECF) predominantly by transporting epithelia in organs such as the gills, intestines,
and kidneys. In fresh water (FW), the ECF of a fish is hyperosmotic compared to its
ambient environment; therefore, a fish must actively absorb ions from water via the gills
and produce large amounts of hypoosmotic urine in the kidney to compensate for passive
ion loss and osmotic water gain (Evans et al., 2005). In sea water (SW), the ECF of a
fish is hypoosmotic compared to its aquatic environment. As a result, SW fish absorb
both salt and water from ingested sea water to promote water absorption in the intestine;
these salts are subsequently excreted actively at the gills, while the kidney produces small
volumes of an isoosmotic urine (Wood and Marshall, 1994).
Fish have varying capacities to expend metabolic resources to fuel active ion
transport (i.e., active salt absorption in FW fish or active ion secretion in marine teleosts)
(George et al., 2006). Furthermore, fish have varying abilities to transform their
transporting epithelia in response to fluctuations in environmental salinity. These
differences in energy reserves and in the capacity for physiological plasticity and
tolerance to osmotic imbalances define the distribution of a fish in its environment. Most
teleost fish tolerate a narrow range of environmental salinities and/or have limited
physiological plasticity. In contrast, euryhaline fish are able to maintain the osmotic
balance of their ECF over a larger range of environmental salinities. This group of fish
can either tolerate osmotic challenges for short periods of time or acclimate to osmotic
1
challenges for prolonged periods while making compensatory adjustments to the
properties of their transporting epithelia.
1.2 Euryhaline Fundulus species – model for environmental salinity tolerance
Some species of fish from the family Fundulus are extremely euryhaline and
either transfer between SW and FW quickly and repeatedly (Evans et al., 2005; Marshall,
2003), or acclimate long term to salinities ranging from FW to several times the strength
SW.
The gulf killifish F. grandis and its sister taxa F. heteroclitus reside in coastal
marshes that are subject to frequent and episodic oscillations in salinity, dissolved oxygen,
and temperature (Marshall, 2003; Wood and LeMoigne, 1991). Their tolerance to
dynamic habitats has made them a widely accepted model for studying the physiological
basis of environmental challenges (Burnett et al., 2007). Although tending to reside in
brackish to marine salinities (Kaneko and Katoh, 2004; Wood and Grosell, 2008; Wood
and Grosell, 2009), killifish are able to tolerate short bouts of freshwater by making rapid,
yet minor physiological adjustments (Marshall, 2003; Wood and Grosell, 2008; Wood
and Grosell, 2009). However, with prolonged exposure to freshwater, F. grandis can
restore ECF osmotic balance by making much larger compensatory adjustments to the
physiology of their ion-transporting epithelia (Wood and Grosell, 2008).
1.3 The influence of environmental salinity on fish gill remodeling
The epithelium that covers the filaments and lamellae of a fish gill separates the
ECF from the external environment (Laurent and Dunel, 1980; Evans et al., 2005). This
gill epithelium is a highly complex, multicellular organ; however, the pavement cells
(PVCs) and the mitochondrion-rich cells (MRCs) are the cell types most often implicated
2
in osmoregulation. PVCs are thin squamous or cuboidal cells with an extensive apical
surface area that are organized in a stratified manner (Wilson and Laurent, 2002). A
special sub-population of PVC, the mitochondrion-rich pavement cells (PV-MRC), exists
in freshwater-acclimated rainbow trout and is characterized by its irregularly-shaped
nucleus, dense peripheral chromatin staining, and absence of a vesicular-tubular network
(Galvez et al., 2002; Goss et al., 2001; Laurent et al., 2006). PV-MRCs are also called
peanut lectin agglutinin (PNA)-negative MRCs because of their abundance of
mitochondria, but their inability to bind peanut-lectin agglutinin (PNA) (Galvez et al.,
2002; Goss et al., 2001; Reid et al., 2003). Although PNA immunoreactivity has not
been described in killifish gills, analogous PV-MRCs in freshwater-acclimated F.
heteroclitus, characterized by their wedge-like appearance at the gill surface between
neighboring PVCs, have been termed cuboidal cells (Laurent et al., 2006).
The ultrastructure and function of the gill epithelium, in particular the structure of
MRCs, are highly variable with environmental salinity. Despite occupying only a
comparably small fraction of the external surface area of the gill, MRCs are essential for
fish osmoregulation. The surface of SW-type MRCs is smooth and concave, and
underlies an apical crypt guarded by PVCs, as shown in Figure 1.1 panel A. The cell
border of MRCs is oval-shaped with poorly developed microvilli towards the center of
cells (Katoh et al., 2001). In contrast, the cell surface of FW-type MRCs is irregularlyshaped, often protruding above the surface of the surrounding PVCs and possessing well
developed microvilli (Katoh et al., 2001; Katoh and Kaneko, 2003), as shown in Figure
1.1 panel B. Most MRCs have a columnar structure that spans the entire thickness of the
epithelium, high mitochondrial density, and abundant expression of transport proteins,
3
making them particularly effective at transepithelial ion movements (Laurent et al., 1995;
Wilson and Laurent, 2002). Though PVCs comprise most of the gill surface, MRCs are
thought to be the primary sites for active ion transport (Evans et al., 2005), and
presumably more susceptible to phenotypic change during environmental stress. MRCs
could physically migrate onto the lamella in order to increase the area for ion uptake after
exposure to soft water (Goss et al., 1998; Perry, 1997; Whitehead et al., 2011).
Moreover, the area of apical surface exposure to the environment and the total number of
MRCs on the gills are altered by disturbances in acid-base balance, or environmental
salinity and oxygen (Claireaux et al., 2004; Dunel-Erb and Laurent, 1980).
Figure 1.1: Scanning electron micrographs on the afferent filament of the gill epithelium
from F. grandis after 3 d exposure to 5 ppt (A) or 0.1 ppt (B). The arrow indicates an
apical crypt of a MRC in a SW-acclimated fish, and the asterisks indicate the apical
surfaces of FW-acclimated MRCs.
The gill epithelium has the ability to remodel itself in response to environmental
and physiological perturbations. This is well exemplified in the crucian carp (Carassius
carassius), which can proliferate its interlamellar cell mass (ILCM) of its gill epithelium
to the extent of completely covering the gill lamellae within hours of exposure to
4
normoxia (Sollid et al., 2003). Although the exact mechanism of gill remodeling remains
uncertain, low oxygen and high temperature both induce gill remodeling by stimulating
apoptosis in the ILCM of crucian carp gill (Sollid and Nilsson, 2006). Moreover,
hypoosmotic conditions stimulate apoptosis or necrosis in F. heteroclitus, although there
is debate regarding the relative importance of each pathway to remodeling (Katoh and
Kaneko, 2003; Laurent et al., 2006). Other stressors, such as heavy metals, also stimulate
cellular remodeling of tilapia (Oreochromis, mossambicus) fish gill by altering the
relative rates of apoptosis and proliferation (Bury et al., 1998).
Cell apoptosis has been widely studied in humans and other mammals systems
(Tran et al., 2004), invertebrate models such as C. elegans (Lettre and Hengartner, 2006),
and fish (Krumschnabel and Podrabsky, 2009). This programmed cell death removes
unwanted or damaged cells in the course of development and differentiation, or in
response to various forms of cellular damage (Taylor et al., 2008). In mammals, there
are two major apoptotic pathways, which include an extrinsic pathway and an intrinsic, or
mitochondrial, pathway. An extrinsic pathway is initiated by ligands, such as tumor
necrosis factor (TNF-α), to form a ligand-receptor complex that activates caspase-8. This
in turn activates caspase-3 to induce cell apoptosis (Tran et al., 2004). The intrinsic
pathway can be stimulated by reactive oxygen species, intracellular stress, and
intracellular damage to activate caspase-3 (Ferri and Kroemer, 2001; Hand and Menze,
2008). Caspase-3, involved in both apoptotic pathways, is a family of highly selective
proteases that cleaves substrates using a cysteine residue (Ferri and Kroemer, 2001; Hand
and Menze, 2008). Although differences exist in the process of apoptosis between fish
and mammals, fish express virtually all of the core components of the mammalian
5
apoptotic machinery (Krumschnabel and Podrabsky, 2009). As the terminal event
preceding cell death, caspase-3 elicits similar effects in both fish and mammals
(Krumschnabel and Podrabsky, 2009; Kuida et al., 1996). Caspase-3, which was the first
caspase identified in teleosts, plays a critical role in development by removing unwanted
cells (Yabu et al., 2001), as well as a central role in the execution-phase of cell apoptosis
induced by different stressors, such as hypoxia, bacterial infection, or hyperthermia in
zebrafish (Lee et al., 2008), Atlantic salmon (Takle et al., 2006), sea bass (Reis et al.,
2007), rainbow trout (Rojo and Gonzalez, 1999), and sturgeon (Lu et al., 2005).
1.4 Transcriptional and enzymatic activities of key enzymes
Although the capacity for gill remodeling to variable environmental salinity is
well reported, few investigations have described the cellular mechanisms that regulatethe
transient and long-term responses of the organ to osmotic challenges. In a recent
transcriptomics study, hypoosmotic shock was shown to alter the expression of 498 genes
(out of approximately 6,800 genes) in the gills of F. heteroclitus during the time course
of acclimation (Whitehead et al., 2011; Whitehead et al., 2012). These transcripts
consisted of the early, transient expression of genes involved in signaling and crisis
control and the altered expression of genes affecting cell morphology, osmoregulation,
and energy metabolism. Transcripts for apoptotic genes, such as cytochrome c, caspase-8,
cyclin-dependent kinase, and p38 MAP kinase, were upregulated upon hypoosmotic
challenge (Whitehead et al., 2011; Whitehead et al., 2012). Additionally, arginase II (arg
II) and ornithine decarboxylase (odc) mRNA levels were among the quickest and most
significantly upregulated by freshwater transfer. These transcripts encode for two
6
enzymes that are involved in polyamine biosynthesis (Whitehead et al., 2011; Whitehead
et al., 2012).
1.4.1 Distribution and function of arginase
Arginase is an enzyme containing a binuclear manganese ion that catalyzes the
hydrolysis of L-arginine to form ornithine and urea (Di Costanzo et al., 2007). It has
been observed in almost all organisms from bacteria (Soru and Zaharia, 1970) and plants
(Muszynsk and Reifer, 1968) to invertebrates (Lisowskamyjak et al., 1978) and
vertebrates (Yu et al., 2003). In most mammals, arginase exists in at least two isoforms,
including arginase I (Arg I) and arginase II (Arg II) (Jenkinson et al., 1996; Spector et al.,
1994). Although the three-dimensional crystalline structures of the two isoforms are
nearly identical, their distribution and function differ (Cama et al., 2003). Arg I is
primarily localized in hepatocytes, and it has been observed in many extrahepatic tissues
such as salivary glands, esophagus, stomach, pancreas, thymus, leukocytes, skin,
preputial gland, uterus, and sympathetic ganglia in mammals (Yu et al., 2003), where it
functions mainly in the ornithine-urea cycle (OUC) (Jenkinson et al., 1996). Arg II is
widely expressed in many extrahepatic tissues, such as intestine and kidney in mammals
(Gotoh et al., 1996). In contrast to extrahepatic Arg I, extrahepatic Arg II has been
implicated in the regulation of arginine and ornithine balance in the venular endothelia
cells of bovine (Li et al., 2001). Arg II also appears to provide ornithine for the
production of putrescine, the smallest of the major polyamines (Jenkinson et al., 1996).
The biological functions of arginase isoenzymes are different in ureotelic and
ammoniotelic species. In ureotelic species, such as mammals, cytosolic arginase I
converts arginine to ornithine, which is shuttled back into mitochondria for incorporation
7
into the OUC (Saha and Ratha, 1989; Jenkinson et al., 1996). In contrast, most waterbreathing organisms, such as teleost fish and other aquatic inverterbrates, are
ammoniotelic, in that they lack a functional OUC and excrete their nitrogenous wastes as
ammonium or ammonia (Hung et al., 2009). In ammoniotelic species, arginase II
catalyzes the reaction converting arginine to ornithine, but do not shuttle this product into
the OUC. Rather, ornithine is metabolized to proline, glutamate, aspartate, glutamine and
polyamines (Gotoh et al., 1996).
Arginase is ubiquitous in different tissues among fish species (Portugal and
Aksnes, 1983). Previous studies have shown that type I and type II arginase (similar
tissue distribution to mammalian Arg I and Arg II) were observed in rainbow trout
(Portugal and Aksnes, 1983; Wright et al., 2004). Moreover, the subcellular localization
of arginase is related to its functions in fish. Mommsen and Walsh (1989) proposed that
hepatic arginase I began to function in the OUC of the lungfish when its subcellular
distribution became cytosolic rather than mitochondrial (Mommsen and Walsh, 1989).
In contrast, coelacanths, marine elasmobranchs, and most teleosts all express Arg II
within hepatic mitochondria. Although it serves no role in the OUC, Arg II plays an
important role in regulating the synthesis of polyamines and proline (Mommsen and
Walsh, 1989; Srivastava and Ratha, 2010). The availability of expressed sequence tag
(EST) for killifish (F. heteroclitus) poses the idea that analogously mammalian Arg I and
Arg II genes may exist in killifish (F. grandis).
1.4.2 Degradation of ornithine decarboxylase
Ornithine decarboxylase (Odc) is a pyridoxal 5’-phosphate-dependent enzyme
that is a homodimer (Tobias and Kahana, 1993). Its active sites are formed at the dimer
8
interface between the NH2-terminal domain of one subunit and the COOH-terminal
domain of another subunit (Bercovich and Kahana, 1993). Odc has a very rapid turnover
rate, with a half-life of less than one hour in brine shrimp and mammals (Svensson et al.,
1997; Watts et al., 1996). One inhibitory protein of Odc is called antizyme, which works
as a non-competitive inhibitor to bind monomer Odc COOH terminus. The binding of
antizyme to the carboxyl terminus of Odc forms an inactive antizyme-Odc complex that
inhibits enzyme activity (Hascilowicz et al., 2002) and exposes its C-terminus to
proteolysis (Bercovich and Kahana, 1993; Coffino, 2001; Heller et al., 1976). Moreover,
α-difluoromethylornithine (DMFO), an irreversible inhibitor of Odc, will inactivate the
enzyme and deplete polyamine reserves upon binding to it (McCann and Pegg, 1992).
DFMO-induced polyamine depletion can impair intestinal epithelial growth
(Bhattacharya et al., 2009), induce intestinal cell apoptosis in mice (Yerushalmi et al.,
2006), and arrest human melanoma cells in the G1 phase (Kramer et al., 2001). At low
concentrations, DFMO depleted intracellular polyamine stores and inhibited cell growth
in human malignant glioma cells (Hunter et al., 1990). DFMO may alter a fundamental
feature of cochlear spiral and cause a hearing loss. DFMO-induced hearing loss may be
mediated through inhibition of polyamine synthesis in cochlea, consequently altering
inward rectification of inner ear-specific Kir currents (Nie et al., 2005). The effects of
DFMO in suppressing the outward component of Kir currents in other systems have been
shown, the functional and clinic significance of DFMO is undetermined (Nichols and
Lopatin, 1997). Moreover, the presence of a DFMO-insensitive decarboxylase was also
detected in brine shrimp in response to altered salt concentrations, and the appearance of
this enzyme may represent an alternative mechanism for polyamine synthesis (Watts et
9
al., 1996). Thus, DFMO is widely used for studying polyamine functions in biological
and medical fields.
1.4.3 Distribution and function of polyamine oxidase
Polyamines are produced in part by the coordinated actions of Arg II and Odc.
Polyamines are a diverse class of aliphatic molecules that are highly polycationic at
physiological intracellular pH (Galston, 1983; Pegg and McCann, 1982; Sekowska et al.,
1998). They have high reactivity to a diverse range of molecules, including DNA, RNA,
and proteins, and perform a large number of physiological functions, such as RNA
transcription, cell growth, cell proliferation, and apoptosis in animals and plants (Galston,
1983; O'Brien et al., 1975; Sekowska et al., 1998). Physiological levels of the
polyamines putrescine, spermidine, and spermine are maintained in cells by both
polyamine biosynthetic and catabolic pathways, as shown in Figure 1.2, and by
regulating oncogenes to form a complex and regulated network, as shown in Figure 1.3.
The activities of Arg II, Odc, SSAT, and Pao contribute to the maintenance of
polyamine concentrations at the physiological level in organisms (Pegg, 1988). Once
Odc converts ornithine to putrescine, this molecule can be further aminated to spermidine
and spermine via the activity of spermidine synthase and spermine synthase, respectively.
Conversely, the polyamine catabolic pathway includes an acetylation reaction by
spermidine/spermine N1-acetyltransferase (SSAT) and the oxidation of N1-acetylspermine
by polyamine oxidase (Pao). Combining SSAT and Pao (SSAT/Pao) can covert
spermine to spermidine, which, in turn, can be converted to putrescine (Figure 1.2).
Although SSAT is a rate-limiting enzyme for polyamine catabolism, Pao is the terminal
oxidase that regulates intracellular polyamine levels (Thomas and Thomas, 2001).
10
Arginine
1
Ornithine
2
Putrescine
6
3
N-Acetylspermine
5
Spermidine
6
4
N-Acetylspermine
5
Spermine
Figure 1.2: Flowchart of the major steps in the mammalian polyamine metabolic pathway
(Wang et al. 2006). Note: 1-arginase II; 2-ornithine decarboxylase; 3-spermidine
synthase; 4-spermidine synthase; 5-spermidine/spermine N1-acetyltransferase; 6polyamine oxidase. Steps 1-4 in blue refer to the biosynthetic pathway, in which
ornithine is converted to putrescine by the action of ornithine decarboxylase. Through a
series of amination reactions, putrescine can be converted to spermidine and spermine.
Steps 5 and 6, shown in red, are part of polyamine catabolic pathway. Through a series of
acetylation and oxidation reaction, spermine and spermidine are converted to spermidine
and putrescine by the action of SSAT and polyamine oxidase.
Pao, a FAD-dependent oxidase that preferentially oxidizes N1-acetylated
polyamine products, was first purified in rat liver (Holtta, 1977). At least two isoforms
of Pao exist in barley (Cervelli et al., 2006), and Pao activity has been observed in rat and
human tissues such as liver, testis, kidney, spleen, small intestine, heart, brain, lung, and
pancreas (Pavlov et al., 1991; Suzuki et al., 1984). Its wide tissue distribution suggests
11
Extracellular signals
Myc
DFMO
Odc
Polyamines
Physiological functions
(e.g., cell growth)
Figure 1.3: The interactions among Myc, ornithine decarboxylase, and polyamines are
shown. Myc genes respond to an extracellular signal and activate Odc translation, which
produces the rate-limiting enzyme for polyamine biosynthesis. Polyamine levels have a
positive feedback on Myc protein expression and form a positive loop. Myc protein
directly participates in multiple physiological functions such as DNA stabilization and
cell growth. α-difluoromethylornithine (DFMO), an inhibitor of Odc, inhibits Myc
protein expression and reduces polyamine levels.
that the regulation of the relative levels of polyamines via modulation of polyamine
catabolism is important in multiple tissues in mammals. Regardless, little is known of
the distribution of Pao activity in teleost fish, nor how it is regulated during osmotic
stress. Pao promotes apoptosis by damaging DNA or lipids (Campestre et al., 2011;
Chaturvedi et al., 2004; Li et al., 2004; Wen et al., 2011), and regulates the ratio of
spermine and spermidine (Adibhatla et al., 2002). Spermine can function as an
antioxidant at high concentrations (Lovaas, 1995) or as a free radical scavenger (Ha et al.,
1998) to benefit cell growth. Moreover, H2O2, which is a by-product of Pao, has
controversial functions in organisms. It can act as an anti-bacterial (Derksen et al., 1999)
12
or fungicidal agent (Marking et al., 1994; Zhang et al., 2004) in fish. In contrast, H2O2
can induce cell apoptosis in mammals by multiple pathways (Chaturvedi et al., 2004;
Demmano et al., 1996; Guan et al., 2011). Therefore, studying the activity of Pao and its
by-product provides further understanding of the polyamine catabolism pathway.
1.5 Physiological function of polyamines to salinity challenge
Environmental challenges such as oscillations in salinity, dissolved oxygen, and
temperature are known to influence polyamine levels in various organisms. Hypersalinity exposure resulted in an increase of putrescine, spermidine, and spermine
concentrations in plants (Campestre et al., 2011), in the gills of the blue crab, C. danae
(Silva et al., 2008), and in the gills of C. sapidus (Lovett and Watts, 1995). Furthermore,
hypoosmotic exposure in brine shrimp increased Odc activity and putrescine
concentration, but had no effect on spermidine and spermine concentrations (Watts et al.,
1996). The collective effect of increasing the mRNA levels of arg II and odc and
reducing the concentration of intracellular polyamine binding proteins during
hypoosmotic shock presumably increased free intracellular polyamine concentrations in
F. heteroclitus (Whitehead et al., 2011), suggesting a possible role of polyamines in the
short-term response to hypoosmotic challenges. Although the role of polyamines during
freshwater challenges is unknown, upregulation of intracellular polyamines or the
exogenous addition of polyamines causes apoptosis, which may be an important
mechanism in gill remodeling (Laurent et al., 2006) or in regulation of cell volume.
Therefore, it is plausible that there exists a close relationship between gill remodeling,
Odc expression, and polyamine levels during hypoosmotic challenges.
13
1.6 Role of polyamines in cell proliferation and cell apoptosis
Polyamines appear to play a dual role in facilitating cell apoptosis and cell
proliferation. Polyamines, the oxidized products of polyamines, and Odc can induce cell
apoptosis, a biological process that is essential for physiological alterations in epithelia
during osmotic challenges (Laurent et al., 2006). Spermine can activate caspases and
initiate apoptosis (Stefanelli et al., 1998; Stefanelli et al., 1999). Moreover, polyamineoxidized products, such as reactive oxygen species (ROS) and polyamine oxidase,
promotes apoptosis by damaging DNA or lipids (Campestre et al., 2011; Chaturvedi et al.,
2004; Li et al., 2004; Wen et al., 2011). Upregulation of spermine oxidase can initiate
apoptosis by oxidative stress or depolarization of mitochondrial membranes (Chaturvedi
et al., 2004). Ornithine decarboxylase can induce apoptosis by triggering the c-Myc
pathway (Packham and Cleveland, 1995; Packham et al., 1996) or can induce spermidine
accumulation to trigger apoptosis (Poulin et al., 1995; Poulin et al., 1989). Inhibition of
ornithine decarboxylase activity with DFMO can inhibit cell apoptosis by depletion of
polyamines (Deng et al., 2005; Monti et al., 1998).
Polyamines are also involved in diverse physiological functions, including the
regulation of cell proliferation and apoptosis. Polyamines are involved not only in
normal cell proliferation and differentiation, but also in malignant transformation. They
can positively regulate proto-oncogenes, such as c-fos, c-ras, and c-myc transcription
(Bachrach et al., 2001; Li et al., 1998; Liu et al., 2005; Tabib and Bachrach, 1994; Tabib
and Bachrach, 1999; Thomas and Thomas, 2003). DFMO application can induce
polyamine depletion and reduce c-myc and c-fos mRNA levels in cells (Celano et al.,
1988; Tabib and Bachrach, 1994; Wang et al., 1993). These findings suggest that high
14
levels of polyamines are associated with many human cancers (Thomas and Thomas,
2003). In addition, polyamines can negatively regulate the expression of growthinhibiting genes, such as p53, and stimulate cell growth (Bhattacharya et al., 2009).
DFMO has been shown to reduce polyamine levels in intestinal epithelia cells by
inhibiting c-myc and c-fos expression (Li et al., 2000) and upregulating p53 protein
expression to hinder cell growth (Bhattacharya et al., 2009; Ray et al., 1999).
Proto-oncogenes increase protein levels, such as c-Myc and c-Fos, via
extracellular stimuli, such as selenium (Yu et al., 2006), oxidative stress (Ichiki et al.,
2003), and salinity (Herbert, 1996); consequently, Odc activity and polyamine
concentrations were increased (Figure 1.3) (Packham et al., 1996). These findings
suggest that a positive feedback loop exists among Myc and/or Fos proteins, Odc, and
polyamines, and that these exhibit physiological impacts as shown in Figure 1.3
(Packham et al., 1996). Although increased Odc activity promotes polyamine
biosynthesis, polyamines have an inhibitory effect on odc mRNA expression (Lovkvist
Wallstrom et al., 2001). Therefore, a complex interplay exists in the cells in order to
support numerous physiological functions of polyamines in animals and plants (Galston,
1983; O'Brien et al., 1975; Sekowska et al., 1998).
1.7 Brief outline of main points of each chapter
Despite recent advances, little is known of the role of polyamines in F. grandis
when faced with hypoosmotic or hyperosmotic challenges. In this dissertation, an
integrative approach, spanning cellular and molecular biology, was utilized to study the
role of polyamines in the short-term and compensatory responses of adult F. grandis to
hypoosmotic challenges.
15
Chapter 2: This chapter aimed to assess the mRNA levels and enzymatic activities
of Arg II and Odc and polyamine concentrations during hypoosmotic challenges. Overall,
my results demonstrated that arg II and odc mRNA levels were highly upregulated in the
gills, with an inverse relationship of salinity on Odc increase during the first few days
post-transfer to hypoosmotic conditions. DFMO inhibited polyamine biosynthesis and
Odc activity following hypoosmotic exposure. Furthermore, polyamine biosynthesis
mediated the early-phase compensatory response of hypoosmotic shock in the euryhaline
Gulf killifish.
Chapter 3: This study aimed to investigate the role of polyamines in gill epithelial
remodeling and opercular epithelial cell swelling. The opercular epithelium is a
surrogate model of the marine fish gill that has been used to study the molecular events
associated with cell swelling and cell volume regulation. Scanning electron microscopy
showed that exposure to fresh water (0.1 ppt) promoted a morphological shift in the
MRCs of the gill epithelium from a SW-type to FW-type. Moreover, addition of
exogenous polyamines, especially spermidine, to the basolateral side of opercular
epithelium during hypotonic exposure, inhibited the decrease of short-circuit current (Isc)
and membrane conductance, suggesting opercular cells may swell by inhibition of Clsecretion. The putative roles of polyamines in regulating cell volume need to be further
studied.
Chapter 4: This study aimed to assess the tissue distribution of key enzymes for
polyamine metabolism. This study demonstrated that hypoosmotic challenges increased
the transcription and enzymatic activities of arginase, ornithine decarboxylase, and
polyamine oxidase in the gills, intestine, and liver. Polyamine metabolism was
16
upregulated in multiple tissues, suggesting that polyamine levels were regulated in the
whole body of the killifish.
17
CHAPTER 2: THE INFLUENCE OF HYPOOSMOTIC EXPOSURE ON POLYAMINE
BIOSYNTHESIS IN THE GULF KILLIFISH
2.1 Introduction
Polyamines are a diverse class of aliphatic, highly reactive molecules due to their
polycationic chemistries at physiological intracellular pH (Galston, 1983; Sekowska et al.,
1998). As such, they have an astonishing array of physiological functions in microbes,
animals, and plants (Galston, 1983; O'Brien et al., 1975; Sekowska et al., 1998),
including their ability to stabilize DNA, RNA, and phospholipids regulate ion channel
activity, facilitate tissue remodeling, and aid in cell growth and differentiation (Pegg and
McCann, 1982; Ray et al., 1999; Tabor and Tabor, 1984). Impairment of this
homeostatic balance can produce severe biological consequences, including nervous
system injury in rat (Adibhatla et al., 2002) and tumor growth in mammal (Thomas and
Thomas, 2003). Due to their reactivity, strict control of polyamine concentrations is
necessary for their physiological functions.
Polyamines are synthesized in part by the coordinated actions of arginase II (Arg
II) and ornithine decarboxylase (Odc). Ornithine is the major precursor of polyamines
and is synthesized from arginine by mitochondrial Arg II. Once produced, ornithine
decarboxylase can convert ornithine to putrescine in an irreversible and rate-limiting
reaction (Pegg and McCann, 1982). Putrescine, which is a diamine, can be converted to
spermidine and spermine, which are triamine and tetramine compounds, respectively.
Once formed, spermidine and spermine can be catabolized back to putrescine in order to
maintain specific ratios of polyamines (Adibhatla et al., 2002). Although under tight
control, the concentrations of these three polyamines are highly affected by various
physiological and environmental stimuli, including osmotic challenges (Zapata et al.,
18
2004), ozone (Langebartels et al., 1991), and UV radiation (Zacchini and de Agazio,
2004) in plants. A recent transcriptomic study on the impacts of hypoosmotic shock in
the euryhaline teleost fish, F. heteroclitus, showed that gill odc mRNA is highly
upregulated almost immediately following acute exposure to fresh water (Whitehead et
al., 2012). It was also observed that the concentration of a gill mRNA expressing a
polyamine-binding protein was decreased to non-detectable levels following freshwater
exposure (0.1 ppt) in F. heteroclitus (Whitehead et al., 2012). Another study observed
that the mRNA level of antizyme, a key inhibitor of Odc activity in vivo, was decreased
in response to hypoosmotic exposure (Mitchell et al., 1998). The collective effect of
increasing polyamine biosynthesis and reducing intracellular polyamine binding would
presumably increase free intracellular polyamine levels. In comparison, the striped
killifish F. majalis, which is unable to tolerate freshwater, showed little upregulation in
polyamine biosynthetic transcripts during hypoosmotic exposure (Whitehead et al., 2013).
These data support a possible role of polyamines in either the short-term or the
compensatory response of these euryhaline fish to hypoosmotic challenges.
The main goal of this study was to assess the influence of acute hypoosmotic
challenges on the mRNA levels and enzymatic activities of Arg II and Odc in the gills of
the Gulf killifish, F. grandis (Hsiao and Meier, 1989). Both Fundulus species are
accepted models for studying the physiological basis of environmental challenges
(Burnett et al., 2007) because they have abilities to tolerate frequent and episodic
oscillations in environmental salinity, dissolved oxygen, and temperature (Marshall,
2003; Wood and LeMoigne, 1991). The gill was investigated because it is one of the
primary organs involved in fish osmoregulation (Evans et al., 2005). This study also
19
investigated the effects of alpha-DL-difluoromethylornithine (DFMO), an irreversible
inhibitor of Odc (McCann and Pegg, 1992), on polyamine regulation during hypoosmotic
challenges in the gill of F. grandis.
2.2 Materials and methods
2.2.1 Experimental animals
Adult F. grandis were obtained from a local hatchery (Gulf Coast Minnows,
Thibodeaux, LA, USA) and held in a 570-L recirculation system maintained in the Life
Sciences Aquatic Facility at Louisiana State University (Baton Rouge, LA, USA). Four
weeks prior to experimentation, fish were randomly transferred to five 110-L aquaria
plumbed on a single recirculation system with mechanical and biological filtration and
UV-sterilization. Water salinity was adjusted by diluting sea salt (Instant Ocean®,
Blacksburg, VA, USA) with reverse osmosis water, and water salinity and temperature
(23 ±1 ˚C) were monitored daily using a water quality meter (YSI, Yellow Spring, OH,
USA). Partial replacement of system water was performed at least twice per week to
keep ammonia and nitrite below the detection levels of commercial kits (Aquarium
pharmaceuticals, Chalfont, PA, USA), and dissolved oxygen was always near airsaturation using a low-pressure aerator. A 12 h light:12 h dark cycle was provided by
fluorescent lighting. Fish were fed a commercial pellet (Cargill, Franklin, LA, USA)
twice daily at 2% body weight per day throughout the study.
2.2.2 Experimental design
In the first series of experiments, one hundred and fifty adult fish (weight range of
4.3-16.8 g) were acclimated to 5 ppt water for at least one month and fed twice daily,
except for 24 h before salinity transfer. Fish were sampled immediately prior to transfer
20
or at 6 h, 1 d, 3 d, or 7 d post-transfer to 5, 2, 1, 0.5, or 0.1 ppt water (N= 6 per time point
per salinity). Fish were net-captured and anesthetized in 0.5 g/L tricaine
methanesulfonate (MS-222). Whole gill baskets were dissected from fish and washed
with ultrapure water for 10 s. The first two left gill arches were immersed in RNAlater
and stored at 4 ˚C overnight, and transferred to -20 ˚C. Total RNA extraction and
quantitative PCR (qPCR) analysis of argII and odc transcript abundance were performed
as described in Section 2.2.3. All four gill arches from the right side were flash frozen in
liquid nitrogen and stored at -80 ˚C. The 1st and 2nd right gill arches were analyzed for
Arg activity, and the 3rd and 4th gill arches were analyzed for Odc activity, as described in
Sections 2.2.4 and 2.2.5, respectively.
In a second series of experiments, the effect of salinity on gill polyamine
concentrations was assessed. Sixty adult fish (weight range of 4.28 to 11.35 g) were
acclimated to 5 ppt water for at least one month, then were sampled randomly prior to
transfer or at 6 h, 1 d, 3 d, or 7 d post-transfer to 5 ppt or 0.1 ppt water (N = 6 per time
point per salinity). Whole gill baskets were dissected from fish, washed with ultrapure
water, flash frozen in liquid nitrogen, and then stored at -80 °C while awaiting analysis of
polyamine concentrations as described in Section 2.2.6.
In addition, the influence of ornithine decarboxylase inhibition on gill polyamine
content in fish undergoing salinity challenges was assessed. Two hundred and forty adult
F. grandis (weight range of 4.9-12.3 g) were acclimated to 5 ppt water. From two days
prior to the start of salinity transfer to one week post-salinity transfer, fish were injected
intraperitoneally once daily at 10 µL/ g with: (i) phosphate-buffered saline (PBS; sham
controls) consisting of 146 mM NaCl, 3 mM KCl, 15 mM NaH2PO4 , 15 mM Na2HPO4,
21
10 mM NaHCO3 (pH 7.4), or (ii) 0.2 mg/µL of alpha-DL-difluoromethylornithine (DFMO)
dissolved in PBS. The dosage of DFMO that affected Odc activity without fish mortality
was determined by pre-experiment (data not shown). Whole gill baskets prior to salinity
transfer (N=6) were sampled. On the third day of injections, half of the PBS-injected fish
(sham control) and half of the DFMO-injected fish were transferred to 0.1 ppt water,
whereas the remaining PBS and DFMO injected fish were maintained at 5 ppt; daily
injections continued for a week post-transfer. In total, fish were allotted to one of four
experimental treatments, including: i) 5 ppt with PBS, ii) 5 ppt with DFMO, iii) 0.1 ppt
with PBS, or iv) 0.1 ppt with DFMO. Whole gill baskets were sampled at 6 h, 1 d, 3 d,
and 7 d post transfer (N=6 per time point per salinity) and stored at -80 °C before
analyzing Odc activity (Section 2.2.5), and polyamine concentrations (Section 2.2.6).
2.2.3 arg II and odc mRNA levels
TRIzol reagent (Invitrogen, Carlsbad, CA, USA) was used for the extraction of
total RNA according to the manufacturer’s instructions. Total RNA concentrations were
determined by measuring the absorbance at 260 nm and 280 nm with a NanoDrop
spectrophotometer (NanoDrop Technologies, Wilmington, DE, USA). The ratio of
absorbance at 260 nm to 280 nm was used to assess the purity of RNA, with a ratio of 1.9
to 2.1 used as the criterion for optimal RNA quality. Single-stranded cDNA was
synthesized from 2 µg total RNA using random primers with a commercial reverse
transcription kit (High Capacity cDNA Reverse Transcription Kit, Applied Biosystems,
Carlsbad, CA, USA).
Expressed sequence tags (EST’s) for arg II (AB290198.1) and odc (CN977303.1)
from F. heteroclitus were obtained from the National Center for Biotechnology
22
Information database (Bethesda, MD, USA). Primers for use in quantitative PCR were
developed for both genes using Integrated DNA Technologies Inc. (Coralville, IA, USA)
software (Table 2.1).
Table 2.1. Sequences of forward and reverse primers used for amplification of F. grandis
arg II, odc, and 18s rRNA amplicons for quantitative RT-PCR analysis.
Name
Direction
Sequence 5’3’
argII-qPCR
Forward
CAGCTGTGGTTACTGCATTGCGTT
argII-qPCR
Reverse
ACAGGAAGCAACAAACCAGCACAG
odc-qPCR
Forward
TTGCCACGCCAATGTGGTCTAA
odc-qPCR
Reverse
ATGCTCCTGTCAAGTAACTGGCCT
18s rRNA-qPCR
Forward
TTCCGATAACGAACGAGAC
18s rRNA-qPCR
Reverse
GACATCTAAGGGCATCACAG
qPCR analysis was performed for the target genes, arg II and odc, and the
reference gene, 18S ribosomal RNA (18s rRNA), using a commercial qPCR kit (SYBR
Green core Reagent, Applied Biosystems, Carlsbad, CA, USA) with an ABI PRISM 7000
Sequence Detection System (Carlsbad, CA, USA). Primer pairs amplified single
amplicons as demonstrated by 2% agarose gel electrophoresis and by denaturation
analysis following individual qPCR runs. Reactions were performed with 0.1 µg firststrand cDNA, 10 nM forward and reverse primers, and 5 µL SYBR Green reagents in a
total volume of 20 µL. Samples underwent 40 PCR cycles at 95 ˚C for 15 s and 60 ˚C for
1 min. Adaptive baselines were used for background detection. The critical threshold
(Ct), the cycle number at which cDNA amplification entered an exponential increase, was
23
calculated automatically using preset algorithms within the software. Preliminary
experiments were performed to calculate qPCR efficiencies for each gene product at
varying concentrations of 1st strand cDNA over a concentration range spanning two
orders of magnitude. One randomly selected control sample (5 ppt pre-transfer control)
was used to develop a standard curve relating Ct to the relative quantity of cDNA. From
this curve, efficiency (E) was calculated from the linear regression of Ct versus log
cDNA for each gene of interest according to the formula:
E = [10(-1/slope)]-1
(Equation 2.1)
Relative change (R) in mRNA levels were measured using the 2 -∆∆Ct method (Fleige et
al., 2006) following the efficiency correction as described by Pfaffl (2001) and is
summarized in Equation 2.2:
R
target
(Equation 2.2)
Target refers to either arg II or odc, and ref refers to the reference gene 18s rRNA.
Efficiency (E) was calculated for each primer set according to Equation 2.1. Reaction
efficiencies for the target and reference genes were between 90 to 110%, with 100%
efficiency representing a doubling in the amount of cDNA per cycle.
Ct represents the
difference in Ct between the target genes (arg II or odc) and the reference gene, 18s
rRNA.
2.2.4 Gill total Arg activity
Total Arg activity was measured from the rate of urea production resulting from
the enzymatic conversion of arginine to ornithine as described by Mommsen (1983). Gill
arches were thawed and immediately homogenized in ice-cold 50 mM HEPES buffer
(V:V for 1:5) using a PRO200 Homogenizer (Lab Depot Inc. Dawsonville, GA, USA).
24
Homogenates were centrifuged at 12,000 g for 4 min at 4 ˚C, then the decanted
supernatants were pre-incubated with 50 mM MnCl2 for 10 min for maximum activation
of the enzyme (Mommsen et al., 1983). Assays were initiated by the addition of 10 µL
of supernatant, 10 µL 50 mM MnCl2, and 480 µL reagent mix, consisting of 50 mM
HEPES (pH 8.0), 250 mM L-arginine, and 1 mM MnCl2. Assays were conducted at room
temperature (23-25 ˚C). Reactions were terminated after 30 min by the addition of 25 µL
70% perchloric acid, then neutralized with KHCO3. Each reaction mix was centrifuged
at 12,000 g for 4 min at 4 ˚C, and the resulting supernatant was analyzed for urea
according to the QuantiChrom urea assay kit instructions (Bioassay System, Hayward,
CA, USA). Total Arg activity was expressed in µmol urea produced /h/g gill.
2.2.5 Gill ornithine decarboxylase activity
Preliminary tests indicated no significant difference in ornithine decarboxylase
enzymatic activity between samples, whether they were flash frozen in liquid nitrogen
and stored at -80 ˚C before analyses or if analyzed fresh following tissue sampling data
not shown). Consequently, frozen gill samples were thawed on ice, homogenized in
reaction buffer (50 mM K2HPO4, 5 mM dithiothreitol, 0.2 mM EDTA, 0.05 mM
pyridoxal 5-phosphate, pH 7.5), and centrifuged at 20,000 g for 20 min at 0 ˚C. The
supernatant was decanted into a 16 mm x 100 mm borosilicate tube sealed with a double
seal rubber stopper (Kimble Chase Kontes, K-882310) and penetrated by a plastic
centerwell (Kimble Chase Kontes, K-882320). The centerwell contained a 1.3 cm2 of
Whatman #1 filter paper saturated with 133 µL 40% KOH. At the start of the assay, a 0.5
µCi mixture of DL-[1-14C] ornithine hydrochloride (52 mCi/mmol, Moravek
Biochemicals, Brea, CA, USA) and cold L-ornithine (Sigma-Aldrich, St. Louis, MO,
25
USA) were added to the air-tight borosilicate tube to reach a final ornithine concentration
of 1.2 mM, as described by Watts (Watts et al., 1996). The reaction mixture was kept in
a water bath at 25 ˚C for 90 min before being terminated with the addition of 5%
trichloroacetic acid.
14
CO2 produced by the enzymatic reaction was collected overnight
on filter paper saturated with 40% KOH. Filter paper was placed into 5 mL Ultima Gold
cocktail (Perkin-Elmer, Waltham, MS, USA), then counted for radioactivity using a
liquid scintillation analyzer (Tri-Carb 2900TR, Perkin-Elmer, Waltham, MS, USA), as
described by Watts (Watts et al., 1996). Internal standardization demonstrated both
constant and negligible quench, so no correction of counting efficiency was necessary.
Odc activity was expressed as nmol CO2 produced /h/g gill.
2.2.6 Gill polyamine contents
Gill baskets were thawed and homogenized with methanol (containing 0.1 mM
HCl) (V:V for 1:3) in dry ice using a PRO200 Homogenizer (Lab depot Inc. Dawsonville,
GA, USA). Samples were incubated in 0.6 M perchloric acid to extract polyamines and
derivatized with o-toluoyl chloride using a slightly modified procedure from that
described previously (Wongyai et al., 1988). Samples (20 µL) were then analyzed for
polyamines by high performance liquid chromatography (HPLC) (Bio-Rad highperformance liquid chromatograph gradient module with a Model 996 photodiode array
detector; Milford, MS, USA). HPLC separations were performed using an ACE C-18PFP column (150 mm x 4.6 mm; 5 µm; Chadds Ford, PA, USA), a mobile phase
consisting of 0.1% trifluoroacetic acid in water and 0.1% trifluoroacetic acid in
acetonitrile, with a flow rate of 1.0 mL/min, and UV detection at 254 nm.
26
2.2.7 Polyamine levels during inhibition of Odc activity in gills
Gill Odc activity and polyamine concentrations of DFMO-injected and PBSinjected fish at 6 h to 7 d post-transfer to 5 ppt or 0.1 ppt water were measured in this
study. The percent change in gill Odc activity and polyamine concentrations of DFMOinjected and PBS-injected fish relative to the simultaneous controls at 6 h to 7 d posttransfer to 5 ppt or 0.1 ppt water, as described in Section 2.2.5, was calculated as shown
in Equation 2.3,
% change
alue post-transfer - alue
alue sham control
100
(Equation 2.3)
where value sham control represents gill Odc activities and polyamine concentrations
measured at 6 h through 7 d post-transfer to 5 ppt or 0.1 ppt water.
2.2.8 Statistical analysis
Statistical differences in gill arg II and odc mRNA levels, enzymatic activity, and
polyamine concentrations between treatments were analyzed using a one-way ANOVA
with a Tukey post hoc test (v.9.3, SAS Institute Inc., Cary, NC, USA). Statistical
differences between treatments, salinity and exposure time were determined using a twoway ANOVA. Statistical differences were considered significant at a P-value of 0.05 or
less.
2.3 Results
2.3.1 Gill arg II and odc mRNA levels
Quantitative PCR analyses of arg II and odc mRNA levels were conducted to
assess the influence of hypoosmotic exposure on the mRNA levels of transcripts
encoding key enzymes for polyamine biosynthesis. The level of arg II mRNA in the gills
27
was inversely correlated with the salinity shortly following acute post-transfer to
hypoosmotic conditions, tending to peak at 1d post-transfer, then gradually decreasing by
7 d post transfer in all hypoosmotic exposures (Figure 2.1 panel A). Gill arg II mRNA
levels were highest in the 0.5 ppt and 0.1 ppt-acclimated fish between 6 h and 1 d
following transfer relative to the pre-transfer controls (P < 0.0001 and P < 0.0001,
respectively) (Figure 2.1 panel A). Gill arg II mRNA in the 0.1 ppt-acclimated fish were
increased 10.5-fold (P < 0.0001) by 1 d post-transfer and remained elevated 2.7-times
above pre-transfer controls by 7 d (Figure 2.1 panel A). Similarly, gill odc mRNA levels
were strongly influenced by the magnitude of hypoosmotic challenges (Figure 2.1 panel
B). Overall, there was a strong dose-dependent decrease in gill odc mRNA levels with
decreasing salinity between 6 h to 7 d (Figure 2.1 panel B). Time and salinity were the
main variables that altered odc mRNA expression (P < 0.0001 and P < 0.0001,
respectively), and there was a statistical interaction between salinity and time (P = 0.0002)
(Figure 2.1 panel B). The gill odc mRNA level increased approximately 21-fold at 6 h
(P = 0.002) and 34-fold at 1 d (P = 0.001) post-transfer to 0.1 ppt water, but was still
elevated approximately 4.5-fold above the level measured in gills of the 5 ppt control
fish by 7 d post-transfer to 0.1 ppt water.
2.3.2 Gill total arginase and Odc activity
Because arg II (Figure 2.1 panel A) and odc mRNA levels were significantly
upregulated (Figure 2.1 panel B) during hypoosmotic exposure, the influence of salinity
on total Arg and Odc activities was investigated. Overall, the magnitude of total Arg
activity was inversely proportional to the magnitude of hypoosmotic shock at 1 d and 7 d
post-transfer (Figure 2.2 panel A) and was only increased by a maximum of
28
Relative arg II mRNA level
14
A
b*
12
10
8
6
4
2
a
0
-2
50
Relative odc mRNA level
0.1 ppt
0.5 ppt
1 ppt
2 ppt
5 ppt
b
b*
b
b
a b
a
a a
0
2
4
6
8
B
40
b*
30
b*
b*
b*
b*
b*
20
b*
b*
b*
10
0
a*
a*
a
a
a
b*
b*
a
a
a
a
b
b
a
a a
a a*
a*
a
a
b*
a
Pre-
-2transfer0
2
4
6
8
Time Post-Transfer (Days)
Figure 2.1: Relative mRNA levels of arg II (A) and odc (B) in gills of F. grandis
acclimated to 5 ppt and sampled pre-transfer or at 6 h, 1 d, 3 d, and 7 d following acute
transfer to 0.1, 0.5, 1, 2, 5 ppt as examined by quantitative RT-PCR analysis. Data are
expressed as mean values ±SEM (N = 6) relative to 18s rRNA after normalizing each
value to the arg II and odc mRNA level of gills from 5 ppt-acclimated fish pre-transfer.
Asterisks indicate a significant difference (P < 0.05) in comparison to the 5 ppt treatment
for that time point; different letters indicate a significant difference (P < 0.05) in
comparison to the pre-treatment for different time points.
29
Arginase activity
(mol urea produced / h / g gill)
380
A
360
b*
340
320
b* b*b*
300
b*
a
a a
b*
a
a
240
4
0
b*
b*
b*
b*
b*
a
a
260
b*
b
280
-2
Odc activity
(nmol CO2 produced / h / g gill)
0.1 ppt
0.5 ppt
1 ppt
2 ppt
5 ppt
2
a
a
4
6
B
b*
3
b*
2
b*
b*
b*
b
1
0
8
a
aa
b a
a
a
Pre-
-2
transfer 0
b*
b
b
a
b
a
a
2
a
4
6
8
Time Post-Transfer (Days)
Figure 2.2: Total Arg activity (A) and Odc activity (B) in gills of F. grandis acclimated to
5 ppt water and sampled prior to transfer, or at 6 h, 1 d, 3 d, and 7 d following acute
transfer to 0.1, 0.5, 1, 2, and 5 ppt water. Data are expressed as mean values ±SEM (N =
6). Asterisks indicate a significant difference (P < 0.05) in comparison to the 5 ppt
treatment; different letters indicate a significant difference (P < 0.05) in comparison to
the pre-treatment.
30
approximately 25% from the pre-transfer value at any given time during exposure (Figure
2.2 panel A). In comparison, gill Odc enzymatic activity increased almost in direct
proportion to the reduction in water salinity at all time points except at 1 d post-transfer
(Figure 2.2 panel B). Time and salinity were the two main variables that affected Odc
enzymatic activity (P < 0.0001 and P < 0.0001, respectively), but there was no statistical
interaction between them (Figure 2.2 panel B). Gill Odc activity was increased
approximately 9.5-fold at 6 h (P < 0.001) and 4.9-fold at 7 d in 0.1 ppt-exposed fish (P <
0.001) compared to 5 ppt-acclimated fish (Figure 2.2 panel B). Odc activity in gills of
0.5 ppt-treated fish were also significantly elevated on 3 d and 7 d post-transfer compared
to the Odc activity of 5 ppt-acclimated fish (Figure 2.2 panel B).
2.3.3. Gill polyamine contents
Based on the large stimulation of the polyamine biosynthetic pathway during
hypoosmotic exposure (0.1 ppt), gill polyamine concentrations were measured. In
accordance with the salinity-dependent increase in gill odc mRNA and Odc activity, the
concentrations of putrescine, spermidine, and spermine in killifish gills were increased
significantly by 0.1 ppt water exposure (Figure 2.3). Time and salinity were the two
main variables to affect gill polyamine levels (P<0.0001 for both effects). Compared to
pre-transfer levels, gill polyamine concentrations did not vary over time in the 5 ppt
treatment; however, gill putrescine level increased after 1 d (P<0.0001) and remained
elevated during the first 7 d post-transfer to fresh water (Figure 2.3 panel A). In
comparison, gill spermidine (Figure 2.3 panel B) and spermine concentrations (Figure 2.3
panel C) were only transiently elevated, decreasing to pre-transfer levels after 3 d of
freshwater exposure.
31
60
55
nmol / g gill
0.1 ppt
5 ppt
A:Putrescine
b*
b*
b*
50
45
40
a
a
35
a
a
30
a
a
25
-2
40
0
4
6
8
B:Spermidine
35
nmol / g gill
2
b*
30
a
b
a
25
a
a
20
a
a
15
a
10
-2
18
0
2
4
6
8
C:Spermine
16
b*
b*
nmol / g gill
14
12
a
10
8
a
a
a
6
a
a
a
4
Pre-
-2
transfer0
2
4
6
8
Time Post-Transfer (Days)
Figure 2.3: Putrescine (A), spermidine (B), and spermine (C) concentrations in gills of F.
grandis acclimated to 5 ppt water then measured pre-transfer, or at 6 h, 1 d, 3 d, and 7 d
post-transfer to 0.1 ppt or 5 ppt water. Data represented as mean values ±SEM (N = 6).
In comparison to the level of 5 ppt treated fish, asterisks indicate a significant difference
in the values of 0.1 ppt treated fish at the same time point (P < 0.05). In comparison to
the pre-transfer level, different letters indicate a significant difference in the values of
post-transfer fish to either 5 ppt or 0.1 ppt at different time points (P < 0.05).
32
The ratios of putrescine level to the sum of concentrations of spermidine and
spermine were summarized in Table 2.2. Hypoosmotic exposure decreased this ratio at
6 h due to the transient accumulation of spermidine and spermine in the gill after acute
salinity challenge (Table 2.2). Over time, spermine and spermidine concentrations
dropped back to 5 ppt levels, whereas gill putrescine concentration remained elevated.
The cumulative effect of these changes in polyamine concentrations over time explain
why polyamine ratios (Table 2.2) increased significantly at 1 d and beyond.
Table 2.2. The ratio of putrescine level over the sum concentration of spermidine and
spermine in the gills of F. grandis for tested salinities over time.
Pre-transfer
6h
1d
3d
7d
[Putrescine] / ([Spermidine]+[Spermine])
5 ppt
0.1 ppt
a
0.86±0.04
---0.94±0.07a
0.81±0.04a,*
0.81±0.14a
0.99±0.11a,*
a
0.82±0.08
1.12±0.16b,*
0.88±0.06a
1.31±0.06b,*
In comparison to the level of 5 ppt treated fish, asterisks indicate a significant
difference in the values of 0.1 ppt treated fish at the same time point (P<0.01). In
comparison to the pre-transfer level, different lower-case letters indicate a significant
difference in the values of post-transfer fish exposed to either 5 ppt or 0.1 ppt at
different time points (P<0.05).
2.3.4 Pharmacological inhibition of ornithine decarboxylase activity
The effects of pharmacological inhibition of Odc activity on polyamine levels in
the gill (Figure 2.4) during hypoosmotic challenge (0.1 ppt) were assessed. Gill Odc
activity was not measured in vivo, but rather the maximal activity that presumably
reflected the amount of active enzyme in the gill. When acclimated to 5 ppt water, gill
Odc activity did not differ between the PBS and DFMO treatments, nor did it vary from
33
the pre-transfer level (Figure 2.4). In contrast, exposure to 0.1 ppt water increased gill
Odc activity significantly at 6 h (39 %) and 1 d (33%) in the sham control fish (Figure
2.4). Following DFMO treatment, Odc activity was reduced by 21% at 1 d and 27% at 3
d significantly compared to the sham control undergoing hypoosmotic transfer (Table
Odc activity (nmol CO2/ h /g gill)
2.3).
2.0
A
b
0.1ppt DFMO
0.1 ppt PBS
5 ppt DFMO
5 ppt PBS
b
1.8
a
a
a
b
1.6
a
a
1.4
a
a
a
1.2
a a
a
a
1.0
0.8
Pre-2
transfer
0
2
a
4
6
8
Figure 2.4: Ornithine decarboxylase activity in gills of F. grandis exposed to 5 ppt and
0.1 ppt water for 7 d and given intraperitoneal injections of PBS or DFMO daily. The
vertical dashed line divides pre-transfer and post-transfer measurements. Data are
presented as mean value ±SEM (N = 6). Asterisks indicate a significant difference
(P<0.05) in comparison to the PBS (sham control) fish; lower-case letters indicate a
significant difference (P<0.05) in comparison to the pre-treated fish.
The concentrations of putrescine, spermidine, and spermine in the gills of 5 pptacclimated fish did not vary over 7 d of sham injection (Figure 2.5). In comparison, gill
polyamine concentrations in fish exposed to 5 ppt water, but injected with DFMO,
decreased significantly (Table 2.3). Putrescine, spermidine, and spermine concentrations
34
were reduced by 86%, 62%, and 47% at 7 d post-transfer to 0.1 ppt water, respectively
(Table 2.3).
Table 2.3. The percent change in gill Odc activity and polyamine concentrations of
DFMO-injected and PBS-injected fish relative to the simultaneous controls at 6 h to 7 d
post-transfer to 5 ppt or 0.1 ppt water.
0.1 ppt
Time
(day)
6h
1d
3d
7d
5 ppt
Odc
PUT
SPD
SP
Odc
PUT
SPD
SP
-15%
-21%*
-27%*
-7%
-37%*
-47%*
-67%*
-75%*
-26%*
-52%*
-24%*
-33%*
-37%*
-47%*
-77%*
-65%*
1%
12%
17%
10%
-36%*
-51%*
-54%*
-86%*
-28%*
-29%*
-24%*
-62%*
-18%*
-21%*
-23%*
-47%*
Asterisks indicate a significant difference (P<0.05) in DFMO-treated fish in comparison
to PBS-treated fish at any given time and salinity. All calculations were calculated using
equation 2.3.
Abbreviations: Odc, ornithine decarboxylase; PUT, putrescine; SPD, spermidine; SP,
spermine.
Compared to pre-transfer levels, the putrescine level in the sham control group
increased over time after transfer to 0.1 ppt water (Figure 2.5 panel A). Furthermore,
DFMO inhibited the increase in putrescine associated with 0.1 ppt exposure at all time
points (Table 2.3). Moreover, the spermidine concentration in gills of 5 ppt-acclimated
fish was significantly reduced over time compared to those of the pre-transfer controls
(Figure 2.5 panel B). Gill spermidine concentration after 1 d of 0.1 ppt exposure was
increased, but returned to the pre-transfer level after 3 d (Figure 2.5 panel B). DFMO
application decreased the spermidine level inducible by hypoosmotic exposure at 1 d and
beyond (Table 2.3). Compared to the pre-transfer level, gill spermine concentration was
only reduced at 7 d in 5 ppt-acclimated fish by DFMO application (Figure 2.5 panel C).
Hypoosmotic exposure increased the spermine level at 1 d, yet had no significant effect
35
after 3 d compared to the pre-transfer levels. (Figure 2.5 panel C). DFMO application
decreased the spermine level inducible by hypoosmotic exposure at 3 d and beyond
Putrescine level (nmol/ g gill)
(Table 2.3).
100
0.1 ppt DFMO
0.1ppt PBS
5 ppt DFMO
5 ppt PBS
A
80
b
b
a
a
a
60
a
a
40
a a
b,c
20
c
b
b
b
c
0
Spermidine level (nmol/ g gill)
-2
60
0
4
6
8
B
50
b
a
40
a
a
a
a
a
30
a
a
a
b
20
b
b
b
c
10
-2
12
Spermine level (nmol/ g gill)
2
0
2
4
6
8
C
10
a
b
a
8
a
6
a a
a
a
4
a
a
b,c
a a
b
2
b
c
0
-2
0
2
4
6
8
Time Post-Transfer (Days)
Figure 2.5: Gill putrescine, spermidine, and spermine concentrations (N=6) in F. grandis
acclimated to 5 ppt and 0.1 ppt water with PBS or DFMO treatment. Data are presented
as mean value ±SEM (N = 6). Lower-case letters indicate a significant difference
(P<0.05) in a pair-wise comparison.
36
2.4 Discussion
2.4.1 Increase in arg II and odc mRNA levels after hypoosmotic exposure
This study suggests that changes in the expression of genes associated with
polyamine biosynthesis may be an important transcriptional response of euryhaline fish
to acute hypoosmotic exposure. This study expands on this knowledge by describing the
magnitude and timing of stimulation of polyamine synthesis, associating it with the early
stages of phenotypic transition of the fish gill to fresh water (data not shown). The
present study suggests that mRNA transcripts of arg II and odc were strongly expressed
in the gills shortly after acute transfer to fresh water (0.1 ppt) and that gill mRNA levels
were inversely correlated to environmental salinity (Figure 2.1 panel A and B).
Moreover, enzymatic activity of Odc was upregulated following acute hypoosmotic
exposure (Figure 2.2 panel B), suggesting a link between hypoosmotic exposure and Odc
activity. A similar relationship between hypoosmotic challenge and Odc activity was
described in the gills of F. heteroclitus (Whitehead et al., 2012) and brine shrimp (Watts
et al., 1996), and in the variant L1210 mouse leukemia cell line (Poulin et al., 1995). As
a key enzyme for polyamine biosynthesis, Odc upregulation suggested a link between
polyamine biosynthesis and hypoosmotic exposure. Although little is known of their
function in fish, polyamines have important roles in cell proliferation and apoptosis (Pegg
and McCann, 1982; Ray et al., 1999; Tabor and Tabor, 1984). Cell proliferation and
apoptosis contributed to the morphological modifaction in F. heteroclitus after
hypoosmotic transfer (Katoh and Kaneko, 2003; Laurent et al., 2006). There is a
possibility that polyamines are involved in morphological modifaction of gills in F.
grandis after hypoosmoitc exposure.
37
Another key enzyme involved in polyamine biosynthesis is arginase, which exists
as at least two different isozymes: arginase I (Arg I) and arginase II (Arg II). Arg I has
been observed in diverse groups of organisms to catalyze the catabolism of arginine to
form ornithine and urea (Wright et al., 2004). Arg I and Arg II have unique functions
based on differential tissue and subcellular distributions. Arg I exists abundantly in the
liver of ureagenic species, where it helps catalyze the reaction of the ornithine urea cycle
(OUC). Arg II is involved in numerous physiological functions including glutamate,
proline, and polyamine biosynthesis, or in the modulation of nitric oxide synthesis in
various organs (Morris, 2004; Li et al., 2001). Although in most mammals, arginase I is
localized to the cytoplasmic compartment, it is expressed in mitochondria of teleost fish
(Wu et al., 1998). However, arginase II is more highly expressed in the mitochondria of
extrahepatic tissues. Most teleost fish are obligate ammonioteles lacking a functional
OUC beyond embryogenesis; thus, Arg II is the major functional arginase in them to
produce ornithine. The current study indicates that the mRNA level and enzymatic
activity of arginase are upregulated in F. grandis during (Figures 2.1 and 2.2). Arginase
activity will constitute the actions of both arginase I and arginase II, despite the fact that
only the latter isoform is likely involved in the formation of ornithine for polyamine
biosynthesis (Wright et al., 2004).
Exposure of F. heteroclitus (Whitehead et al., 2011; Whitehead et al., 2013) and
F. grandis to fresh water has been shown to strongly upregulate gill ornithine
decarboxylase mRNA levels. This study demonstrates that odc mRNA and Odc activity
were highly inversely correlated with water salinity, and that these effects occurred
concomitant with an increase in gill polyamines. In contrast, the marine species
38
Fundulus majalis, which is intolerant to fresh water, exhibits limited capacity to
upregulate odc mRNA levels during hypoosmotic exposures (Whitehead et al. 2013).
This pattern suggested that the transcriptional changes in key enzymes of polyamine
biosynthesis corresponded to the osmoregulatory abilities within Fundulus species.
Moreover, the difference in polyamine level is likely to contribute to species-specific
osmoregulatory variation in Fundulus (Whitehead et al., 2011; Whitehead et al., 2013).
The roles of polyamines may be related to regulation of cell volume and tissue plasticity
in euryhaline fish.
2.4.2 Effects of hypoosmotic exposure on gill polyamine levels
This study suggests that hypoosmotic exposure stimulates polyamine biosynthetic
pathways, resulting in an increase in putrescine, spermidine and spermine concentrations
in the gill (Figure 2.3). Similar results have been found in brine shrimp (Watts et al.,
1996) and mammalian cells (Munro et al., 1975). The relative abundance of the different
classes of polyamines may result in the differences in physiological functions during
osmosis challenge. Putrescine, spermidine, and spermine concentrations increased in
plants (Campestre et al., 2011) and in the gills of the crab Callinectes danae (Silva et al.,
2008) with hyperosmotic exposure. Moreover, putrescine and spermidine concentrations
in the gills of Callinectes sapidus were also elevated after exposure to full-strength sea
water (Lovett and Watts, 1995). In contrast, putrescine concentrations in Artemia nauplii
(Watts and Good, 1994) and in Eschericia coli (Munro et al., 1972) decreased with an
increase in environmental salinity. Only putrescine level was increased in brine shrimp
during hypoosmotic exposure (Watts et al., 1996). Although their functions are not clear,
polyamines on one hand are protetive of cellular function but on the other hand are
39
cytotoxic. Thus, regulation of polyamine homeostasis in the face of environmental
stressors such as osmotic challenges is paramount. Moreover, the rapidity of changes in
polyamine concentrations may play an important role in cell volume regulation because
they are responsible for initial efflux of intracellular inorganic ions and intracellular
organic compounds (Watts et al., 1996). Further research is necessary to elucidate the
specific relationship between polyamines and cell volume regulation. The high ratio of
the level of putrescine to the sum levels of spermidine and spermine (Table 2.2)
suggested polyamine catabolism regulated the ratio of different polyamine components,
and resulted in putrescine accumulation (Adibhatla et al., 2002). Although the role of
putrescine during hypoosmotic challenges has not yet been determined, previous studies
have shown that putrescine regulates protein transcription (Rhee et al., 2007) and
remodels microtubules in mammals (Mechulam et al., 2009). Thus, it is plausible to
suggest that putrescine is involved in the gill remodeling, but further studies are needed.
2.4.3 Effects of DFMO on polyamine contents during the hypoosmotic exposure
DFMO is a cancer chemopreventive agent that is used for the prevention of
various types of cancer due to its inhibition of polyamine synthesis (Nie et al., 2005), but
the physiological effects of DFMO other than on Odc activity are uncertain. DFMO can
covalently bind and inactivate Odc (McCann and Pegg, 1992), and thus reduce polyamine
levels (Hunter et al., 1990; Bhattacharya et al., 2009). This study showed that the
efficacy of DFMO on Odc activity was dependent on exposure salinity. For instance,
DFMO significantly reduced the ability of freshwater exposure to stimulate Odc activity
in killifish gills, leading to a concomitant reduction in polyamine levels in the tissue
(Table 2.3). In comparision, DFMO had no effect on endogenous Odc activity in fish
40
gills in the absence hypoosmotic exposure (Table 2.3). Surprisingly, gills still exhibited a
marked reduction in gill polyamine concentrations with DFMO despite the inability to
inhibit the enzyme. DFMO-sensitive decarbolxylase was detected in IEC-6 cells
(Bhattacharya et al., 2009) in human melanoma cells (Kramer et al., 2001) and human
malignant glioma cells (Hunter et al., 1990). Moreover, the presence of a DFMOinsensitive decarboxylase was detected in brine shrimp in response to altered salt
concentrations (Watts et al., 1996). DFMO-insensitive decarboxylase may lack critical
residues for binding DFMO that limit the ability for inhibition by DFMO (Jhingran et al.,
2003). Although the reason for this discrepancy is unclear, there are two possible
explanations for this reduction of polyamine despite no inhibition of Odc activity. These
changes in Odc activity are more likely due to the changes occurring in cellular
polyamine contents when DFMO is administered. Polyamines exert an inhibitory effect
on Odc synthesis, and previous studies have shown that putrescine, spermidine, and
spermine inhibited Odc activity to 85%, 46%, and 0% of the levels in the control group,
respectively, in intestinal epithelial crypt (IEC-6) cells (Iwami et al., 1990). Moreover,
polyamines stimulated Odc degradation by inducing the synthesis of antizyme (Heby and
Persson, 1990; Svensson and Persson, 1996). In this study, intracellular polyamine levels
are much lower in 5 ppt-acclimated fish than 0.1-acclimated fish after DFMO treatment
(data not shown). The polyamine-mediated feedback regulation of Odc is attenuated in 5
ppt -acclimated fish; thus, Odc activity may keep the original level in 5 ppt -acclimated
fish. Another explanation is the existence of DFMO-insensitive decarboxylase.
Although there is no direct evidence of a DFMO-insensitive Odc in the gills, the relative
percentages of DFMO-sensitive Odc and DFMO-insensitive Odc can be made by looking
41
at the total Odc activity and the amount inhibited by DFMO. The insensitivity towards
DFMO may due to substitution of key substrate binding residues in active site pocket
(Preeti et al., 2013). The activity of DFMO-insenstitive Odc is one component of total
Odc in the gills, but its activity was not affected by DFMO application. However, the
data in the current study are insufficient to know the function of DFMO-insensitive
decarboxylase during osmotic challenge. The existence of DFMO-insensitive
decarboxylase may be involved in the general catabolic reduction of the intracellular
amino acid pool that occurs at low salinity (Watts et al., 1996). Moreover, its occurrence
may represent an alternative pathway for polyamine biosythensis (Watts et al., 1996).
Further studies are necessary to determine the nature and role of this enzyme.
Additional studies are needed to explore the role of polyamines during the initial
shock to and recovery from hypoosmotic exposure in F. grandis. Previous results
suggest that there may be a close relationship between Odc expression, polyamines levels
and osmotic challenges, although the existence of DFMO-insensitive Odc in killifish
complicates this relationship. Although the mechanism is unclear, polyamines may be
involved in gill remodeling and acclimation to environmental salinity fluctuation.
However, the fact that putrescine, not spermidine or spermine, increased after prolonged
freshwater exposure suggests that polyamine catabolic pathways may be upregulated over
time due to hypoosmotic shock.
42
CHAPTER 3: THE POTENTIAL ROLE OF POLYAMINES IN GILL EPITHELIAL
REMODELING DURING EXTREME HYPOOSMOTIC CHALLENGES IN THE
GULF KILLIFISH
3.1 Introduction
The Gulf killifish, F. grandis, and its sister taxa, F. heteroclitus, reside in coastal
marshes that are subject to frequent and episodic oscillations in salinity, dissolved oxygen,
and temperature (Marshall, 2003; Wood and LeMoigne, 1991). Their ability to tolerate
these dynamic habitats has made them a widely accepted model for studying the
physiological basis of tolerance to environmental stressors (Burnett et al., 2007).
Although they prefer brackish to marine salinities (Kaneko and Katoh, 2004; Wood and
Grosell, 2008; Wood and Grosell, 2009), euryhaline Fundulus species can tolerate
reduced environmental salinity and can take short-term forays into fresh water for feeding
(Marshall, 2003). Killifish appear to cope with short bouts of fresh water by making
rapid, yet easily reversible physiological adjustments (Marshall, 2003; Wood and Grosell,
2008; Wood and Grosell, 2009); only with prolonged exposure to fresh water are they
required to make more substantial compensatory adjustments to the physiology of their
ion-transporting epithelia (Wood and Grosell, 2008).
One of the most important organs in fish osmoregulation is the gill, which
consists of several cell types that mediate the active excretion of salts in marine fish and
the active uptake of osmolytes in freshwater fish (Chang et al., 2001; Evans, 2011; Evans
et al., 2005). It is generally accepted that the gill epithelium has the ability to remodel
itself in response to environmental and physiological perturbations; however, few studies
have delineated the salinity at which the gill epithelium transitions from a seawater to a
freshwater morphology (Laurent et al., 2006; Katoh et al., 2001; Katoh and Kaneko,
2003). Most Fundulus species are marine teleosts that have the ability to retain their
43
seawater physiology at salinities approaching fresh water (Feldmeth and Waggoner, 1972;
Griffith, 1974; Whitehead, 2010). Copeland and Philpott suggested that F. heteroclitus
made this physiological transition at a salinity between 2 ppt and 0.2 ppt based on a
morphological assessment of the fish gill surface (Copeland, 1950; Philpott and Copeland,
1963). Wood and Grosell (2009) reported that the transepithelial potential, an important
determinant of the electrochemical driving force of gill ion transport, reversed in polarity
instantaneously following acute transfer of seawater-acclimated F. heteroclitus to fresh
water, but that only with prolonged exposure to fresh water did the gills of F. heteroclitus
functionally “switch” into one characteristic of a freshwater fish (Wood and Grosell,
2008; Wood and Grosell, 2009). Recently, it has been shown that F. heteroclitus
acclimated to sea water could be transferred abruptly to 0.4 ppt with no significant effects
to their plasma osmolyte concentrations over 14 d post-transfer (Whitehead et al., 2012).
Although the ability of the gill to remodel with varying environmental salinities is well
reported, few investigations have described the cellular mechanisms regulating the
transient and long term responses of the fish gill to osmotic challenges.
It has recently been shown that the upregulation of transcripts encoding enzymes
involved in polyamine biosynthesis is an early compensatory response to hypoosmotic
exposure in the F. heteroclitus gill (Whitehead et al., 2011). Polyamines are a family of
organic cations in cells, and their accumulation are affected by environmental salinity in
bacteria (Peter et al., 1978), plants (Zapata et al., 2004) and mammalian cells (Poulin et
al., 1991). Polyamines, which include the multi-aminated molecules, putrescine,
spermidine, and spermine, increase in concentration in the killifish gill during
hypoosmotic challenge (0.1 ppt), a response associated with increased transcription and
44
enzymatic activity of ornithine decarboxylase, the rate-limiting step that converts
ornithine to putrescine (Chapter 2). Following 3 d of acute transfer from sea water to
fresh water, only gill putrescine levels remained upregulated in killifish, suggesting a
distinct role of this polyamine in freshwater acclimation or in the maintenance of the
freshwater gill phenotype; still, little is known of the physiological function of these
molecules in fish.
The main goal of this study was to characterize the role of polyamines in
osmoregulation, gill remodeling, and apoptosis in the Gulf killifish, Fundulus grandis,
during exposure to a highly-resolved range of low environmental salinities. This salinity
range was chosen because it was known to demarcate the transition between the marine
and freshwater gill osmoregulatory phenotypes (Feldmeth and Waggoner, 1972; Griffith,
1974; Whitehead, 2010; Marshall, 2003). The effects of alpha-DLdifluoromethylornithine (DFMO), an irreversible inhibitor of ornithine decarboxylase, on
plasma chemistry and the expression of various apoptotic markers in the fish gill during
freshwater transfer were studied. Finally, the effects of polyamine supplementation on
active ion transport and membrane conductance in the opercular epithelium under
hypoosmotic conditions were studied. The opercular epithelium has been used as a
surrogate model of the marine fish gill (Zadunaisky et al., 1995; Zadunaisky, 1996 ) to
study the mechanisms of cell volume regulation during osmotic challenges (Marshall et
al., 2005).
45
3.2 Materials and methods
3.2.1 Experimental animals
Adult F. grandis were obtained from a local hatchery (Gulf Coast Minnows,
Thibodeaux, LA) and kept in the Life Sciences Aquatic Facility at Louisiana State
University (Baton Rouge, LA, USA). Fish were maintained in 5 ppt water for at least
one month in a 570-liter glass aquarium with mechanical and biological filtration, and
UV-sterilization. Water salinity was monitored using a water quality meter (YSI, Yellow
Spring, OH, USA) and temperatures were kept at 22-24 ˚C. Fish were kept on a light
cycle of 12 h light and 12 h dark provided by fluorescent lighting set on an automatic
timer. Fish were fed a commercial fish pellet Cargill, Aquaxcell™ WW Fish Starter
4512, Franklin, LA, USA) twice daily at 2% body weight throughout experimentation,
and partial water changes were performed at least twice per week to keep water quality
constant.
3.2.2 Experimental protocols
Series 1: Blood plasma chemistry and gill surface morphology were assessed
during acute hypoosmotic challenges. One hundred and fifty adult fish (weight range
4.3-16.8 g) were acclimated to 5 ppt water for at least one month. Immediately prior to
sampling, a subset of fish (N=6) were sampled as described below. The remaining fish
were divided randomly into 5 groups, then transferred acutely to 5, 2, 1, 0.5, or 0.1 ppt
water, which was made by diluting Instant Ocean® salt (Blacksburg, VA, USA) with
reverse osmosis water. At 6 h, 1 d, 3 d, or 7 d post-transfer (N = 6 per salinity per time
point), fish were net-captured and anesthetized with 0.5 g/L tricaine methanesulfonate
(MS-222). Whole gill baskets were dissected from fish and washed with MilliQ-purified
46
water for 10 s. The 3rd and 4th left gill arches were fixed in a solution of 2%
glutaraldehyde and 1% formaldehyde in 0.1 M cacodylate buffer for scanning electron
microscopic (SEM) analysis of the gill surface, as described in Section 3.2.3. Blood was
collected into micro-hematocrit capillary tubes (Fisher Scientific, Pittsburgh, PA, USA)
following caudal severance and centrifuged at 3,000 g for 3 min. Plasma was collected
and stored at -20 ˚C while awaiting analysis of plasma osmolality, as described in Section
3.2.4.
Series 2: Previous studies have shown that the mRNA abundance and enzymatic
activity of ornithine decarboxylase is significantly upregulated in gills of F. grandis posttransfer to 0.1 ppt water (Chapter 2). It has also been established that inhibition of
ornithine decarboxylase during hypoosmotic challenge significantly reduces the
concentrations of all three major polyamines in the gill (Chapter 2). In these studies, the
influence of inhibition of ornithine decarboxylase on plasma osmolality, plasma Na+ and
Cl- concentrations, and caspase-3 activity were assessed during hypoosmotic exposure.
One hundred and twenty adult F. grandis (weight range 4.9-12.3 g) were acclimated to 5
ppt water, and fish were weighed and injected intraperitoneally with 10 µL phosphatebuffered saline (PBS; sham control; 146 mM NaCl, 3 mM KCl, 15 mM NaH2PO4, 15
mM Na2HPO4, and 10 mM NaHCO3 at pH 7.4) or DFMO (0.2 mg DFMO/µL PBS) per g
fish for two days prior to hypoosmotic transfers. At 3 d, half of the fish in the DFMO
and PBS treatments were transferred to 0.1 ppt water and the other half of the remaining
fish were transferred to 5 ppt water. Fish were injected daily with either DFMO or PBS,
as described above. Plasma was collected using the same procedure as described for
Section 3.2.2 (series 1) and stored at -20 oC while awaiting analysis of plasma chemistry,
47
as described in Section 3.2.4. All four gill arches from the right side were flash frozen in
liquid nitrogen and stored at -80 ˚C for analysis of caspase-3 activity, as described in
Section 3.2.6.
Series 3: Considering the extensive remodeling of the gill epithelium (Figure 3.2)
and the significant increase in transcripts encoding enzymes involved in polyamine
biosynthesis (arginase II and ornithine decarboxylase) (Chapter 2), the effects of salinity
challenges on the mRNA levels of arginase II (arg II) and ornithine decarboxylase (odc)
in isolated mitochondrion-rich cells (MRCs) and pavement cells (PVCs) of the gill
epithelium were investigated. Forty-eight fish (weight range 8.19-17.83 g) were
acclimated in 5 ppt water for at least one month. Half the fish were transferred to 5 ppt
water and the other twenty-four fish were transferred to 0.1 ppt water. After one day
post-transfer, fish were sacrificed and their gills removed. MRCs and PVCs were
isolated from pooled fish gills (i.e., 2 gills per isolation) using the procedure described in
Galvez et al., 2002, with modifications described below. Briefly, F. grandis fish gills
were washed three-times in ice-cold phosphate-buffered saline (137 mM NaCl, 4.3 mM
Na2HPO4, 2.7 mM KCl, and 1.4 mM NaH2PO4 at pH 7. 8) for 10 min. Gill filaments
were removed from their arches, cut into small sections, and digested in 0.25% TrypsinEDTA (Sigma, St. Louis, MO, USA) for 20 min at 300 RPM using a SK-300 shaker (Lab
companion, Geumcheon-Gu, Seoul, Korea). Gill digests were passed through a 96 µm
mesh filter in 15 mL stop buffer (Kelly et al., 2000). Gills were trypsin-digested 2 or 3
times to yield a mixed population of isolated gill cells, then centrifuged at 300 g for 8 min
at 4 ˚C. Gill cells were diluted in a 10-fold volume of PBS centrifuged at 500 g for 10
min at 4 ˚C. The pellet, which was resuspended in 2 mL PBS, was applied to the top of a
48
discontinuous Percoll gradient (Sigma, St. Louis, MO, USA) consisting of a 1.03 g/ml
top layer, a 1.06 g/ml middle layer, and a 1.09 g/ml bottom layer. The stacked cell
suspension was centrifuged (1,000 g for 35 min at 4 °C) to isolate PVCs and MRCs at the
1.03-1.06 g/mL and 1.06-1.09 g/mL Percoll interfaces, respectively. Each isolated cell
fraction was stored at -80 °C until RNA extraction, then analyzed by quantitative PCR for
arg II and odc mRNA levels, as described in Section 3.2.5.
Series 4: Tests were conducted to assess the putative role of polyamines in the
compensatory response of fish to osmotic shock. The opercular epithelium was used as a
surrogate of the fish gill to study the influence of polyamines on membrane conductance
(Gt, mS cm-2) and short-circuit current (Isc, µA cm-2) (Marshall et al., 2000). Adult F.
grandis were acclimated to 5 ppt water and anaesthetized in 0.2 g/L MS 222. Both
opercular membranes of the fish were isolated as described by Marshall et al (2000).
Briefly, each operculum was mounted between two plexiglass sliders (Physiological
instruments, San Diego, CA, USA), exposing a 0.1 cm2 diameter round aperture, and
placed in individual Ussing chambers (Physiological instruments, San Diego, CA, USA)
(Marshall et al., 2000). The Ussing chambers contained isotonic saline (151 mM NaCl, 3
mM KCl, 0.88 mM MgSO4, 4.6 mM Na2HPO4, 0.48 mM K2HPO4, 1.0 mM CaCl2, 11.0
mM HEPES-free salt, and 4.5 mM urea at an osmolality of 334 mOsm/kg and pH 7.8) on
each side of the membrane, as described previously (Daborn et al., 2001). The isotonic
bathing solution was bubbled with 100% oxygen. The basolateral side of the membrane
was bathed in 2 mL isotonic saline with 3 mM glucose added, whereas the apical side
contained 2 mL isotonic saline. Isc and Gt across the opercular epithelium were measured
in real time with a data acquisition system (PowerLab and LabChart 7 software,
49
ADInstrument Inc., CO, USA). A pulse of 0.002 V was given every min to calculate Gt
according to Ohm’s law: Gt= Isc / V. Once a baseline was achieved under symmetrical
conditions (isotonic saline on both sides of the membrane), the isotonic solution was
replaced with a hypotonic bathing solution (75% isotonic saline, 25% distilled water)
under symmetrical conditions, resulting in an osmolality of 270 Osm/kg. This hypotonic
solution mimicked the plasma osmolality of killifish during the early stages of
hypoosmotic shock.
Isc and Gt were assessed in opercula immersed in isotonic (Iso) and hypotonic
(Hypo) bathing solutions for 30-45 min each to allow steady-state values to be reached.
Preliminary experiments were conducted to assess the influence of polyamine
concentration on Isc during hypotonic shock. Briefly, baseline Isc for opercular epithelium
exposed to Iso and Hypo solutions were determined. Individual polyamines at final
concentrations of 0.4-400 mM putrescine, 0.3-300 mM spermidine, and 0.1-100 mM
spermine (N=6 per concentration per polyamine) were added to the basolateral side of
opercular epithelium (V:V of 1:1000) and Isc was recorded until it reached a new steady
state (approximately 25-30 min). These data were used to derive the relationship
between changes in Isc versus polyamine concentrations for each of the three compounds
(data not shown). Since there was no significant difference in change in Isc change
between low dose and high dose of individual polyamines, a polyamine mixture
consisting of 0.4 mM putrescine, 0.3 mM spermidine, and 0.1 mM spermine was used in
subsequent experiments as described below. First, the influence of polyamine mixture on
the change of Isc and Gt was assessed. Briefly, a steady state of Isc and Gt was attained
under Iso condition, then a polyamine mixture (0.4 mM putrescine, 0.3 mM spermidine,
50
0.1 mM spermine) was added to the basolateral side of opercular epithelia, either a)
coincident with transfer from isotonic to hypotonic conditions (pre-PA, N=7), or b)
following initial hypotonic transfer and establishment of a steady-state Isc under
hypotonic conditions (post-PA, N=10). Second, the influence of individual polyamines
on the change of Isc and Gt was assessed. 0.4 mM putrescine (PUT), 0.3 mM spermidine
(SPD), and 0.1 mM spermine (SP) were added to the basolateral side of opercular
epithelia coincident with transfer from isotonic to hypotonic conditions, respectively
(N=7 for Hypo+PUT, Hypo +SPD and Hypo +SP). Isc and Gt were recorded for 40 min
until reaching a steady-state, and percent change of Isc and Gt was measured according to
Equation 3.1,
Percent change
alue treat - alue isotonic
alue isotonic
(Equation 3.1)
where Value treat represents the values of Isc or Gt at pre-PA, post-PA, Hypo, Hypo+PUT,
Hypo +SPD and Hypo +SP at different time points. Value isotonic indicates a steady-state
of Isc or Gt when opercular epithelia were incubated in isotonic saline.
Series 5: Killifish gill c-fos and c-myc mRNA transcript levels and caspase-3
activity were assessed as putative measures of cell proliferation and cell apoptosis
following acute freshwater transfer. Sixty adult fish (weight range 3.83-12.61 g) were
acclimated to 5 ppt water for at least one month. Gills were sampled prior to transfer and
at 6 h, 1 d, 3 d, and 7 d post-transfer to 5 ppt or 0.1 ppt water (N = 6 per salinity per time
point). The first two left gill arches were immersed in RNAlater and stored at 4 ˚C
overnight, then transferred to -20 ˚C while awaiting total RNA extraction and qPCR
analyses of c-fos and c-myc transcript abundance, as described in Section 3.2.5. All four
51
gill arches from the right side were flash frozen in liquid nitrogen, stored at -80 ˚C, and
used for analysis of caspase-3 activity, as described in Section 3.2.6.
3.2.3 Scanning electron microscopy (SEM) analyses
Gills were fixed in 2% glutaraldehyde and 1% formaldehyde in 0.1 M cacodylate
for 4 h, rinsed in 0.1 M cacodylate buffer containing 0.02 M glycine, post-fixed in 2%
osmium tetroxide for 1 h, and rinsed in water. Samples were dehydrated in an ethanol
series (10 min at 50%, 10 min at 70%, 10 min at 80%, 2 rinses of 10 min at 95%, and 3
rinses of 15 min at 100% ethanol), dried with liquid CO2 in a Denton critical point dryer
(Cherry Hill, NJ,USA,) mounted on aluminum SEM stubs and coated with a
gold:palladium solution (V:V of 60:40) in an Edwards S150 sputter coater (Ashbord,
Kent, UK). The afferent edge of each gill filament was randomly imaged using a JSM6610 high vacuum mode SEM (Tokyo, Japan).
3.2.4 Plasma chemistry analyses
Plasma osmolality was analyzed by freeze-point depression (Precision systems
INC, Natick, MA, USA). Plasma Na+ concentrations were measured using flame atomic
absorption spectroscopy (Varian Australia Pty Ltd, Australia), and plasma Clconcentrations were measured using a modified mercuric thiocyanate method (Zall et al.,
1956).
3.2.5 Relative arg II, odc, c-myc, and c-fos mRNA levels
Commercial SYBR Green master mix (QIAGEN Inc. Valencia, CA, USA) was
used to analyze the relative mRNA levels of arg II, odc, c-myc, c-fos, and the reference
gene, 18 S ribosomal RNA (18s rRNA). Expressed sequence tag (EST) sequences for arg
II (AB290198.1), odc (CN977303.1), c-fos (DN956552.1) and c-myc (CN985702.1) from
52
F. heteroclitus were obtained from the National Center for Biotechnology Information
database (http://www.ncbi.nlm.nih.gov/). Primers for quantitative PCR were developed
for all genes using Integrated DNA Technologies Inc. software (Coralville, IA, USA)
(Table 3.1). A 20 µL reaction mixture (0.1 µg cDNA, 10 nM forward and reverse
primers, 5 µL SYBR Green reagents) was used for the ABI Prism 7000 SDS system
(Carlsbad, CA, USA). The first strand of cDNA was reverse-transcribed from total
mRNA of MRCs or PVCs isolated from the fish gill. Samples were cycled 40 times at 95
˚C for 15 s and 60 ˚C for 1 min. Efficiency and relative changes in mRNA levels were
measuring using the 2 -∆∆Ct method (Fleige et al., 2006).
Table 3.1. Sequences of forward and reverse primers used for quantitative RT-PCR
analyses of c-myc, c-fos, arg II, odc, and 18s rRNA amplicons in F. grandis gill.
Sequence 5’3’
Name
Direction
c-myc
Forward
TGCTTGTGCCTCTCACCAGTTCTA
c-myc
Reverse
AGCCTTCGAATCCTCCACGTTCTT
c-fos
Forward
ATCTGACAGCATCAAGTGCCTCCT
c-fos
Reverse
AGGTCTGGACGTTGCAGACTTCAT
argII
Forward
CAGCTGTGGTTACTGCATTGCGTT
argII
Reverse
ACAGGAAGCAACAAACCAGCACAG
Odc
Forward
TTGCCACGCCAATGTGGTCTAA
odc
Reverse
ATGCTCCTGTCAAGTAACTGGCCT
18s rRNA
Forward
TTCCGATAACGAACGAGAC
18s rRNA
Reverse
GACATCTAAGGGCATCACAG
53
3.2.6 Caspase-3 activity assay
A caspase-3 assay kit (Sigma, St. Louis, MO, USA) was employed to measure
caspase-3 activity, which is based on the hydrolysis of acetyl Asp-Glu-Val-Asp 7-amido4-methylcoumarin by caspase 3 that results in the release of the fluorescent 7-amino-4methylcoumarin (AMC). Gill arches were homogenized with a 10-fold volume of 0.85%
NaCl, and the original homogenate was used directly to measure caspase-3 activity.
Original gill homogenates were diluted 20-fold and measured for total protein (Sigma, St.
Louis, MO, USA). Fluorescence intensity was measured with excitation at 355 nm and
emission at 460 nm using a VICTORTM X3 multilabel plate reader (Perkin Elmer, Santa
Clara, CA, USA). A 1x assay buffer of 200 µl served as a blank to adjust the fluorimeter
to zero. Fold–change in gill caspase-3 activity was calculatedby taking the ratio of the
enzymatic activity of post-transfer to 5 ppt water or 0.1 ppt water over the enzymatic
activity of pre-transfer at each time point.
3.2.7 Statistical analysis
Statistical differences in plasma osmolality, gene transcripts, and Isc and Gt
between treatments were analyzed using a one-way ANOVA (v.9.3, SAS Institute Inc.,
Cary, NC, USA). Statistical differences were considered significant at a P-value of 0.05
or less.
3.3 Results
3.3.1 Plasma chemistry
Initial experiments were performed to assess the influence of comparably small
changes in environmental salinity on plasma osmolality (Figure 3.1) and gill surface
morphology (Figure 3.2). The magnitude of the reduction of plasma osmolality
54
disturbances depended on the severity of the hypoosmotic transfer and on the time
following post-transfer. Plasma osmolality decreased by approximately 19.5% following
transfer from 5 ppt water to either 0.1 ppt or 0.5 ppt water, reaching their lowest values at
1 d and 6 h, respectively (Figure 3.1). Plasma osmolality in the 0.5 ppt treatment
recovered to its original level by 1 d, but it did not completely recover to the pre-transfer
value even after 7 d in the 0.1 ppt treatment (P=0.0069). In comparison, plasma
osmolality in F. grandis was not affected by the abrupt transfer from 5 ppt to 1 ppt, 2 ppt,
or 5 ppt water at any time during the 7 d post-transfer period.
440
Plasma osmolality (mOsM /Kg)
a
420
a
a a
a
400
380
360
a
a
a
a
a
a
a
a
a
a
b
b
b*
0.1 ppt
0.5 ppt
1 ppt
2 ppt
5 ppt
340
320
a
b* b*
300 Pre-2transfer 0
2
4
6
8
Time Post-Transfer (Days)
Figure 3.1: Plasma osmolality (N = 6) in F. grandis acclimated to 5 ppt water then
sampled at pre-transfer or at 6 h, 1 d, 3 d, and 7 d following acute transfer to 0.1, 0.5, 1, 2,
and 5 ppt water. The vertical dashed line divides pre-transfer and post-transfer time
points. Data represent the mean ±SEM, mOsM/kg. Asterisks indicate a significant
difference (P ≤ 0.05) in comparison to the 5 ppt treatment for that time point; different
letters indicate a significant difference (P ≤ 0.05) in comparison to the pre-transfer for a
given salinity.
55
3.3.2 Gill morphology
Scanning electron microscopy was used to assess the morphology of the apical
opening of MRCs on the gill surface of fish undergoing osmotic shock, as described in
Section 3.2.3. The gill surface of fish acclimated to salinities of 1-5 ppt water consisted
of pavement cells (PVCs) possessing concentric rings of microridges around the outer
cell margins, and, on occasion, microvilli projecting towards the center of cells (Figure
3.2).
6h
1d
3d
7d
5 ppt
2 ppt
1 ppt
0.5 ppt
0.1 ppt
Figure 3.2: Representative scanning electron micrographs of the afferent filamental
epithelium of F. grandis gills after acclimation to 5 ppt and acute transfer to 0.1, 0.5, 1, 2,
or 5 ppt water at 6 h, 1 d, 3 d, and 7 d post-transfer. Asterisks indicate pavement cells,
solid white arrows point to seawater-type mitochondrion-rich cells (MRCs), and dashed
white arrows point to freshwater-type MRCs. Scale bar 5 μm.
56
MRCs in fish gills exposed to 5 and 2 ppt water were predominately seawatertype, as noted by their small apical openings at the gill surface which were localized to
the afferent edge of the filamental epithelium. Exposure to 1 ppt water led to minor
changes in the surface appearance of the fish gill, with a slight enlargement of MRCs
during the initial 6 h to 1d post-transfer. FW-type MRCs, characterized by their convex
shape and well-developed microvilli, began to emerge on the gill surface as salinity
decreased to 0.5 ppt and below. These cell types were only expressed transiently, being
replaced by SW-type MRCs, which reemerged by 3 d at 0.5 ppt or by 7 d at 0.1 ppt.
Regardless, FW-type MRCs with smaller apical openings were still present following 7 d
exposure to 0.1 and 0.5 ppt water (Figure 3.2).
3.3.3 argII and odc mRNA levels in pavement cells and mitochondrion-rich cells
Considering the significant increase in arg II and odc mRNA levels in the gill
during freshwater acclimation (Chapter 2), the cellular distribution of these increases in
transcript abundance was assessed. Compared to the PVCs at 5 ppt, the levels of arg II
and odc mRNA in MRCs did not change. In contrast, arg II and odc mRNA levels
increased by 11.4-fold (P=0.0037) and 8.1-fold (P=0.0009) in MRCs upon transfer of
fish to 0.1 ppt water (Figure 3.3). During hypoosmotic exposure, arg II (Figure 3.3 panel
A) and odc (Figure 3.3 panel B) mRNA levels were 4.9-fold (P=0.0224) and 2.6-fold
(P=0.0113) higher in MRCs relative to PVCs, respectively.
3.3.4 Plasma chemistry during ornithine decarboxylase inhibition
Compared to the pre-transfer level, plasma osmolality declined approximately 20%
after 1 d hypoosmotic transfer in the sham control (P=0.0039). In contrast, fish
57
10
10
A
b*
B
b*
8
6
4
a
a
Relatvie odc mRNA
Relative argII mRNA
8
MRCs
PVCs
2
6
a
4
2
a
a
a
0
0
5 ppt
0.1 ppt
5 ppt
0.1 ppt
Figure 3.3: Relative arg II and odc mRNA levels in mitochondrion-rich cells (MRCs) and
pavement cells (PVCs) in gills of fish acclimated to 5 ppt, then acutely transferred to 5
ppt or 0.1 ppt water for 1 d. The mRNA levels were normalized to the mean value of
expression in PVCs from 5 ppt-acclimated fish. Data represent the mean ± SEM (N=6).
Different letters indicate a significant difference (P≤0.05) in comparison to the 5 ppt
treatment for the same cell type; asterisks indicate a significant difference (P≤0.05) in
comparison to PVCs for that salinity.
administered DFMO, showed no effect of freshwater transfer over time (Figure 3.4 panel
A). Plasma osmolality was not significantly different between the DFMO-treated fish
and PBS-treated fish maintained in 5 ppt water over time (Figure 3.4 panel A). Plasma
Na+ concentrations decreased by 14% at 6 h and 18% at 1 d following transfer to 0.1 ppt
water hypoosmotic exposure in sham controls, but returned to the pre-transfer level by 3
d (Figure 3.4 panel B). The plasma Na+ loss induced within the first day of by freshwater
exposure (0.1 ppt) was attenuated by administration of DFMO (Figure 3.4 panel B),
although it to returned to the pre-transfer level by 3 d. Compared to the pre-transfer level,
plasma Cl- concentration decreased at 6 h and 1 d following hypoosmotic exposure in
sham controls. DFMO application decreased the reduction of plasma Cl- induced by
hypoosmotic exposure at 6 h and 1 d (P=0.0084 and 0.016, respectively) (Figure 3.4
58
panel C). Both plasma Na+ and Cl- returned to pre-transfer levels after 3 d of
hypoosmotic exposure in shams. Moreover, the concentrations of plasma Na+ and Cl- in
DFMO and PBS-treated fish at 5 ppt water were not significantly different over time
Plasma osmolality (mOsm/kg)
(Figure 3.4 panel B and C).
450
a
a
400
a
a
a
a
a
a
300
a
a
a
a
a
250
180
a
a
a
350
b*
-2
Plasma Na+ concentration(mM)
0.1 ppt DFMO
0.1 ppt PBS
5 ppt DFMO
5 ppt PBS
A
0
2
B
a
160
6
a
a
a
4
8
a
a
a
140
ba
a
a
a
a
a
a
b* b*
120
100
80
-2
Plamsa Cl- concentration (mM)
180
0
2
4
6
8
C
160
140
120
a
a a
a
a
100
a
a
a*
a*
a
a
a
a*
a
b*
80
b*
Pre-2
transfer
0
2
4
6
8
Time post-transfer (Days)
Figure 3.4: Plasma osmolality (mOsM/kg) (A), plasma sodium concentration (mM) (B),
and plasma chloride concentration (mM) (C) in F. grandis acclimated to 5 ppt and
sampled at pre-transfer or at 6 h, 1 d, 3 d, and 7 d following acute transfer to 0.1 ppt or 5
ppt water. Fish received intraperitoneal injections of DFMO or PBS (sham control) (see
text for details). The vertical dashed line divides pre-transfer and post-transfer. Data
represents the mean ±SEM (N=6). Asterisks indicate a significant difference (P≤0.05) in
comparison to the 5 ppt PBS treatment for that time point; letters indicate a significant
difference (P≤0.05) in comparison to the pre-treatment for a given salinity.
59
3.3.5 The effect of polyamines on opercular Isc and Gt
The influence of polyamines on the hypotonic-induced changes in Isc and Gt in the
opercular epithelium was investigated. Isc of the opercular epithelium dropped
immediately after transfer to the hypotonic solution, reaching a new steady state within
30-35 min (Figure 3.5 panel A). Hypotonic shock induced a 52% reduction in Isc
compared to that measured in the isotonic medium (Figure 3.5 panel A). Compared to
the hypotonic control, pre-PA significantly inhibited Isc reduction in the opercular
epithelium after 20 min hypotonic exposure, reaching its maximum inhibition at 40 min
(P=0.0006) (Figure 3.5 panel A). However, post-PA did not affect Isc compared to the
hypotonic control over time (Figure 3.5 panel A). Hypotonic shock immediately
decreased Gt, which could not be inhibited by either pre-PA or post-PA application over
time (Figure 3.5 panel B).
Additional experiments were conducted to assess the contribution of individual
polyamines to changes in Isc and Gt during hypotonic shock. Isc and Gt of the opercular
epithelium dropped immediately after transfer to the hypotonic solution, reaching new
steady states within 30-35 min (Figure 3.6 panel A). Compared to the hypotonic control
(52% decrease), only spermidine inhibited the reduction of Isc and Gt over time, having
its maximal effect at 40 min (P<0.01) (Figure 3.6). Hypotonic shock significantly
decreased Gt, which was not inhibited by the application of putrescine or spermine
(Figure 3.6 panel B).
3.3.6 Relative c-myc, c-fos mRNA expression, and caspase-3 activity
The mRNA levels for c-fos and c-myc were measured in the gills following
hypoosmotic exposure as a measure of cell proliferation in the tissue. Gill c-fos mRNA
60
A Iso
Hypotonic
Control
pre-PA
post-PA
Percent change of Isc
0
-20
*
*
*
20
30
40
30
40
-40
-60
0
10
B Iso
Hypotonic
Percent change of Gt
0
-20
-40
-60
0
10
20
Time (min)
Figure 3.5: Percent change (relative to baseline isotonic conditions) in the short circuit
current, Isc (A) and membrane conductance, Gt (B) of opercular epithelia kept in
symmetrical conditions to isotonic (Iso) (N=10) or hypotonic (Hypo) solutions (N=10).
The vertical dashed line represents the transfer between these two media. Controls
represent those epithelia receiving no administration of the polyamine mixture. Asterisks
represent significant differences from hypotonic control exposures at the same time
points (P<0.05). A polyamine mixture (0.4 mM putrescine, 0.3 mM spermidine, and 0.1
mM spermine) was added to the basolateral side of opercular epithelium, either a)
coincident with transfer from isotonic to hypotonic conditions (pre-PA), or b) following
initial hypotonic transfer and establishment of a steady-state Isc under hypotonic
conditions (post-PA).
61
A Iso
Hypotonic
Control
+PUT
+SPD
+SP
0
Percent change of Isc
-10
-20
*
-30
*
*
*
20
30
40
**
**
30
40
-40
-50
-60
-70
-10
0
10
B Iso
Hypotonic
Percent change of Gt
0
-10
-20
-30
*
*
10
20
-40
-50
-60
-10
0
Time (mins)
Figure 3.6: Percent change (relative to baseline isotonic conditions) in short circuit
current, Isc (A) and membrane conductance, Gt (B) of the opercular epithelium in isotonic
(Iso) or hypotonic (Hypo) solutions. The vertical dashed line represents the transfer
between these two media. Data represents the mean ±SEM (N=6). Controls represent
those epithelia receiving no addition of polyamines. Single asterisks represent significant
differences from hypotonic challenges at the same time points (P≤0.05); double asterisks
represent significant differences from hypotonic challenges at the same time points
(P≤0.01).
62
expression increased 14-fold at 6 h (P<0.0001) and 4-fold at 1 d (P=0.001) post-transfer
to 0.1 ppt water, but returned to the pre-transfer level by 3 d (Figure 3.7 panel A).
Compared to the 5 ppt control, the c-fos mRNA level increased 10.5-fold and 4.4-fold at
6 h and 1 d, respectively, following hypoosmotic exposure (P<0.02 for both time points)
(Figure 3.7 panel A). Gill c-myc mRNA abundance was increased 6.1-fold after 6 h of
hypoosmotic exposure, but returned to the pre-transfer level at 7 d (Figure 3.7 panel B).
The level of c-myc mRNA increased up to 7.1-fold by 1 d post-transfer (compared to the
pre-transfer level) (P<0.0001) (Figure 3.7 panel B). c-myc mRNA expression increased
at 6 h, 1 d, and 3 d in the gills of 0.1 ppt-acclimated fish (P=0.001, 0.011, and 0.029,
respectively) (Figure 3.7 panel B).
Based on the gill remodeling observed in Figure 3.2, the onset of apoptosis was
assessed by analyzing caspase-3 activity in gill. Compared to the pre-transfer caspase3activity, caspase-3 activity in the gill did not change significantly during exposure to 5
ppt water (Figure 3.7 panel C). However, gill caspase-3 activity increased approximately
3-fold following 3 d of hypoosmotic exposure (P=0.0062) and returned to the control
level at 7 d (Figure 3.7 panel C). Compared to 5 ppt, hypoosmotic exposure (0.1 ppt)
increased gill caspase-3 activity significantly at 1 d (P=0.0429) and 3 d (P=0.0399)
(Figure 3.7 panel C). After hypoosmotic exposure, gill caspase-3 activity was increased
at 6 h and 1 d significantly (P<0.0001for both time points) compared to the sham control
(Figure 3.7 panel D), and this upregulation was inhibited by 38% at 6 h (P=0.0002) and
43% at 1 d (P<0.0001) after administering DFMO (Figure 3.7 panel D).
63
A
Relative c-myc mRNA expression
b*
15
10
b*
5
a
a
0
a
a
Pre-
-2transfer 0
Fold change of Caspase-3 activity
20
0.1 ppt
5 ppt
4
a
a
a
2
4
6
C
b*
a
1
a
0
Pre-
-2transfer 0
a
a
a
4
6
a
0
a a
a
8
a
a
a
2
4
D
b*
2
6
b*
a*
a
a
a
1
a
a
a
a
a
Pre-
-2transfer 0
8
0.1 ppt DFMO
0.1 ppt PBS
5 ppt DFMO
5 ppt PBS
3
0
2
b*
b*
5
4
b*
a
10
Pre-
3
a
0.1 ppt
5 ppt
-2transfer 0
0.1 ppt
5 ppt
2
B
15
8
Fold-change of Caspase-3 activity
Relative c-fos mRNA expression
20
2
a
a
a
a
4
6
8
Time Post-Transfer (Days)
Figure 3.7: Relative c-fos (A) and c-myc (B) mRNA levels (relative to the pre-transfer
levels), and caspase-3 activity without DFMO application (C) and with DFMO
application (D). The vertical dashed line divides pre-transfer and post-transfer. Data
represent the mean ±SEM (N=6). Asterisks indicate significant differences (P≤0.05) in
comparison to the 5 ppt PBS treatment for that time point; letters indicate a significant
difference (P≤0.05) in comparison to the pre-treatment for different time points.
3.4 Discussion
3.4.1 Gill epithelium morphology
In the present study Gulf killifish were acclimated to 5 ppt water (~15 % sea
water), then subjected fish to small reductions to environmental salinity. This study
highlighted the extent to which only a small reduction in environmental salinity (i.e.,
64
from 1 ppt to 0.5 ppt) produced a dramatic change in plasma osmotic balance and gill
surface morphology. Plasma osmolality was significantly reduced at 6 h post-transfer to
0.5 ppt water, although it recovered back to the 5 ppt control level within 1 d (Figure 3.1).
Fish transferred to 0.1 ppt water experienced a similar reduction in plasma osmolality
(Figure 3.1), although osmotic balance was not restored to pre-transfer levels even after
14 d exposure (data not shown). The results showed that the apical surface of the gill
MRCs of F. grandis retained an apical crypt characteristic of a seawater fish gill at
salinities of 1 ppt and above (Figure 3.2). Only when acutely exposed to salinities at or
below 0.5 ppt did the apical MRCs on the fish gill surface attain a concave appearance
more characteristic of a freshwater gill phenotype (Figure 3.2). In fact, gill surface
morphology, as determined using SEM (Figure 3.2), was shown to change from its basal
SW-type phenotype only in treatments and at time points in which fish had also
experienced reductions in plasma osmolality (Figure 3.1). The drastic decrease of plasma
osmolality during hypoosmotic exposure was considered to be the result of diffusional
ion loss through the body surfaces. When plasma osmolality decreased, the size of
MRCs increased as shown in Figure 3.2. It has been demonstrated that MRCs alternate
their morphology to meet abrupt environmental osmotic change (Kaneko and Katoh,
2004). Fish exposed to salinities of 0.1 ppt or 0.5 ppt exhibited few seawater-type MRCs
after 6 h; these cells were replaced by large MRCs that were predominantly triangular in
shape, with apical surfaces that contained short microvilli flush with the surrounding
PVCs (Figure 3.2). Laurent (2006) termed these freshwater-type MRCs as cuboidal cells,
characterized by their wedge-like appearance between neighboring PVCs, analogous to
the mitochondrion-rich PVCs described in rainbow trout (i.e., PNA- MRCs; see Chapter
65
1). With continued extreme hypoosmotic exposure, the surface exposure of these
cuboidal cells decreased over time and seawater-type apical crypts emerged, even in fish
acclimated to 0.1 ppt water (Figure 3.2). This produced a mixed population of SW and
FW MRCs, as previously described (Laurent et al., 2006; Scott et al., 2006; Scott et al.,
2008 ; Whitehead et al., 2012). It is unclear why SW MRCs began to resurface in the 0.1
ppt- and 0.5 ppt-acclimated fish. Laurent et al. (2006) suggest that seawater MRCs either
undergo apoptosis, necrosis, exfoliation, or are simply covered by pavement cells.
SEM images showed a transition between the two morphologies, in which
freshwater-acclimated Fundulus lacked the apical cavity characteristics of seawateracclimated fish (Copeland, 1950; Kaneko and Katoh, 2004; Katoh et al., 2001; Katoh et
al., 2000; Laurent et al., 2006). This ability to rapidly transition based on environmental
salinity is beneficial for euryhaline fish species, such as F. grandis. One possible
mechanism may be related to the regulation the Cl- ion transport across the gill epithelium.
In order to acclimate to fresh water, killifish must actively absorb ions such as Na+ and
Cl- from dilute aquatic environment or inhibit the loss of plasma Na+ and Cl- to
compensate for diffusive ion loses and osmotic water gain. The freshwater-acclimated
fish had cuboidal cells (as observed by scanning electron microscopy) that intercalated
between PVCs or other MRCs forming a relatively impermeable barrier to ions (Kaneko
et al., 2002; Katoh and Kaneko, 2003), preventing ion loss in the gill epithelium. The
morphological transition was also accompanied by the loss of apical chloride channels
(cystic fibrosis transmembrane conductance regulator), leading to an inhibition of Clsecretion (Katoh et al., 2001; Katoh and Kaneko, 2003). In summary, the appearance of
66
cuboidal cells presumably would help inhibit ion loss at the gill epithelium and maintain
plasma ion concentrations in response to rapid fluctuations in environmental salinity.
3.4.2 Possible roles of polyamines in cell apoptosis and cell proliferation
The regulation of cell apoptosis and caspase-3 activity may be related to gill
remodeling during compensation to salinity challenges. The present study demonstrated
that relative arg II and odc mRNA expression was elevated in the MRCs of 5 pptacclimated fish gills and that these transcripts further increased in abundance in this cell
type upon 0.1 ppt water exposure (Figure 3.3). Arginase II is abundant in extrahepatic
cells rich in mitochondria, where it may be involved in numerous physiological functions
including glutamate, proline, and polyamine biosynthesis, or in the modulation of nitric
oxide synthesis in various organs (Morris, 2004). The concurrence of increasing arg II
and Odc mRNA levels in MRCs (Figure 3.3) and MRC remodeling (Figure 3.2)
suggested that polyamines may be associated with the phenotypic plasticity of MRCs in
the fish gill. Although the role of polyamines in MRCs remodeling of killifish is unclear,
polyamines can cause cell apoptosis (Gerner and Meyskens, 2004), which is thought to
mediate gill remodeling in killifish (Laurent et al., 2006). Moreover, hypoosmotic
challenges led to a significantly transient increase in gill putrescine, spermidine and
spermine concentrations (Chapter 2). Polyamine accumulation occurred concomitantly
with an increase in gill caspase-3 activity following 0.1 ppt water exposure (Figure 3.7
panel C). Caspase-3 activity also increased in zebrafish during DNA damage (Lee et al.,
2008), in Atlantic salmon (Salmo salar) during hyperthermia (Takle et al., 2006), in sea
bass (Dicentrarchus labrax L.) during bacterial infection (Reis et al., 2007), in rainbow
trout after hyperosmotic transfer (Rojo and Gonzalez, 1999), and in sturgeon (Acipenser
67
schrenckii) during prolonged hypoxia (Lu et al., 2005). The role of caspase-3 in fish may
correspond with extensive apoptosis and morphological abnormalities in tissues
(Yamashita et al., 2008). The results in the present study indicated that polyamines may
activate the intrinsic apoptotic pathway (chapter 1) via caspase-3, as seen in previous
studies (Poulin et al., 1995; Stefanelli et al., 1998; Tobias and Kahana, 1993).
Additionally, accumulation of caspase-3 during hypoosmotic challenge may be involved
in the regulation of tissue plasticity (Snigdha et al., 2012).
The effect of DFMO on caspase-3 activity in the fish gill during freshwater
transfer was studied. DFMO application resulted in the reduction of gill polyamine levels
during hypoosmotic exposure (Chapter 2). Hypoosmotic challenge increased caspase-3
activity at 6 h and 1 d (Figure 3.7 panel D); however, applying DFMO decreased gill
caspase-3 activity, as observed in intestinal epithelia (Deng et al., 2005; Ray et al., 2000),
in bone marrow stromal cells (Muscari et al., 2005) and cardiac myoblasts (Tantini et al.,
2006). DFMO application resulted in decreasing of both polyamine levels and caspase-3
activity, suggesting a link betwewn polyamine levels and cell apoptosis (Figure 3.7).
This reaction benefits morphological transitions in the gill epithelium, because seawater
MRCs must undergo apoptosis to reduce the number of seawater MRCs, or they can be
replaced by freshwater MRCs (Laurent et al., 2006). Therefore, it is plausible to suggest
a link between polyamine levels, cell apoptosis and gill remodeling.
Recent results have shown that increased ornithine decarboxylase activity and
polyamine levels at the 6 h and 1 d (Chapter 2) were concomitant with c-myc and c-fos
transcription increases in killifish gills (Figure 3.7 panel A and B). This relationship was
described in a previous study in rat kidney cells (Tabib and Bachrach, 1994). Although
68
the relationship between increasing polyamine levels and transcriptional regulation of cfos and c-myc genes in the gill after hypoosmotic challenge exists, the causal relationship
between them is unclear. Previous studies have shown that polyamines, including
putrescine, spermindine and spermine, can positively regulate c-fos, c-ras, and c-myc
transcription (Bachrach et al., 2001; Li et al., 1998; Liu et al., 2005; Tabib and Bachrach,
1994; Tabib and Bachrach, 1999; Thomas and Thomas, 2003). Moreover, c-myc and cfos mRNA expression was inhibited by DFMO, an inhibitor of ornithine decarboxylase,
in carcinoma cells (Celano et al., 1988; Wang et al., 1993) and in cultured cells (Tabib
and Bachrach, 1994). These studies suggest that polyamines have the ability to stimulate
transcriptional c-fos and c-myc genes during environmental challenges (Yuen et al.,
2001).
3.4.3 The influence of polyamines on osmotic properties in the opercular epithelium
Most eukaryotic cells have a polyamine transporter system on their cell
membrane that facilitates the internalization of exogenous polyamines (Wang et al.,
2003). Intracellular polyamines can plug the receptor channel pore, or permeate the ion
channel of these receptors. Therefore, the interactions of intracellular polyamines and ion
channels have been reported (Williams, 1997). Exogenous polyamines modulate some
types of glutamate receptors, such as increase the size of glutamate receptor (Williams,
1997). Overexpression of intracellular polyamine or exogenous addition of polyamines
causes the loss of intracellular polyamine homeostasis (Gerner and Meyskens, 2004).
Recently, the interactions of intracellular polyamines with different types of ion channels
have been reported, including blocking of some types of ion channels and glutamate
receptors (Williams, 1997). Polyamines are believed to regulate cell volume during
69
hypotonic challenges (Watts et al., 1996). Following transfer from 5 ppt to either 0.1 ppt
or 0.5 ppt, the plasma osmolality dropped approximately 20% at 1 d and 6 h post transfer,
and required approximately 3 d to return to its original level (Figure 3.1). Zadunaisky et
al. (2005) suggested a relationship between plasma osmolality and Cl- secretion rate in
MRCs. In order to test this hypothesis, the opercular epithelium is used as a surrogate of
marine fish gill. The opercular epithelium of killifish, a gill-like membrane rich in MRCs,
is used to study the mechanism of active Cl- secretion in vitro. When the opercular
epithelium was subjected to hypotonic solutions on the basolateral side, the Isc
dramatically decreased (Figure 3.5). In this tissue, Isc is almost entirely derived a Clsecretory current, a distinguishing feature of a marine phenotype (Degnan et al., 1977;
Krogh, 1938). Isc decreased immediately across the opercular membrane upon hypotonic
exposure, which could be explained partly by swelling-activated Cl- channels on the
basolateral membrane (Hoffmann et al., 2002; Hoffmann et al., 2007 ).
Pre-treatment with a polyamine mixture containing all three polyamines or
spermidine alone decreased the magnitude of the hypotonically-induced reduction in Isc
immediately (Figure 3.5 panel A), which indicates that polyamines moderate the
inhibition of active Cl- secretion in vitro. The paradox is that this effect prevents the
reduction in epithelial Cl- efflux associated with hypotonically induced cell swelling.
Moreover, exhausting polyamine levels by DFMO (Figure 3.4 panel C) attenuated the
decrease in Cl- transport across the epithelium during hypoosmotic exposure, which
indicates that polyamines prevented the reduction in Isc associated with cell swelling. It
is unclear about the mechanism; however, endogenous polyamines are known to block
ion channels, such as inward rectifier K+ channels (Doupnik et al., 1995), intracellular K+
70
and Ca2+ channels (Scott et al., 1993), and Na+ and Ca2+ fluxes (Bowie and Mayer, 1995;
Isa et al., 1995), to control the resting membrane potential and the excitability of cell
membranes. Polyamines also moderate Na+/K+/2Cl- co-transporter (NKCC) and cystic
fibrosis transmembrane conductance regulator (CFTR), which are involved in mediating
cell swelling during hypotonic shock (Hoffmann, 1992; Hoffmann, 2000; Hoffmann et al.,
2002; Hoffmann et al., 2007 ).
The overall conductance, as a rough estimate of ionic permeability, is lower in
euryhaline fish acclimated to hypotonic media compared to the level in isotonic media
(Figure 3.5 panel B and 3.6 panel B), as shown in previous studies (Marshall et al., 2000;
Motais et al., 1969). Spermidine pretreatment prevented both a decrease in membrane
conductance (Gt) (Figure 3.6 panel B) and Isc (Figure 3.6 panel A). The gill epithelium is
more permeable to cations than anions (House and Maetz, 1974), which may explain why
Cl- transport was inhibited by spermidine over time (Figure 3.6 panel B). Although the
mechanism is currently unknown, polyamines, especially spermidine, inhibit the
reductions in Isc and Gt. These findings may help maintain Cl- concentration and regulate
the amount of seawater-type MRCs in gill epithelia.
Polyamines may facilitate cell apoptosis and cell proliferation with the goal of
remodeling the gill, especially in MRCs. The remodeling of MRCs is a rapid
physiological adjustment to cope with short bouts of freshwater exposure, and helps
euryhaline fish survive in environmental salinity perturbations. However, polyamine
accumulation inhibited Cl- secretion from MRCs, and resulted in MRC swelling. Further
studies are needed to determine the role of polyamines in cell volume regulation and gill
remodeling.
71
CHAPTER 4: THE INFLUENCE OF OSMOTIC STRESS ON TRANSCRIPTION AND
TRANSLATION OF ARGINASE, ORNITHINE DECARBOXYLASE, AND
POLYAMINE OXIDASE IN TISSUES
4.1 Introduction
Previous studies have shown that hypoosmotic exposure can stimulate polyamine
biosynthetic pathways, leading to an increase in putrescine, spermidine, and spermine
concentrations in the killifish gill (Chapter 2). With prolonged fresh water exposure,
spermidine and spermine concentrations decrease to control levels, whereas the gill
putrescine concentration remains elevated (Chapter 2). These data suggest the polyamine
biosynthetic pathway is upregulated during acute hypoosmotic transfer and that the
polyamine catabolic pathway may become upregulated for late compensatory response of
hypoosmotic transfer.
Polyamines are produced in part by the coordinated actions of arginase II (Arg II)
and ornithine decarboxylase (Odc) (Pegg and McCann, 1982). Arginase catalyzes a
reaction converting arginine to ornithine, a substrate of ornithine decarboxylase (Odc).
Odc is a rate-limiting enzyme for the production of polyamines (Pegg and McCann,
1982), catalyzing the reaction converting ornithine to putrescine. This is the first step in
the polyamine biosynthetic pathway (Tobias and Kahana, 1993). Putrescine is converted
to spermidine and spermine via spermidine synthase and spermine synthase, two enzymes
that had low turnover rate and are important sites for polyamine biosynthesis. The
polyamine catabolic pathway consists of reactions involving the acetylation and oxidation
of larger polyamines into samller polyamines. It primarily includes the acetylation of
spermidine or spermine by spermidine/spermine -N1-acetyltransferase (SSAT) and the
oxidation of N1-acetylspermidine and N1-acetylspermine to putrescine and spermidine,
respectively by polyamine oxidase (Pao) (Seiler, 2004). Though both SSAT and Pao are
72
key enzymes for polyamine catabolism, Pao is the terminal oxidase that regulates the
ratio of spermidine to spermine (Adibhatla et al., 2002).
The key enzymes for polyamine metabolism have wide tissue distribution.
Previous studies have shown that Arg II is widely expressed in many extrahepatic tissues,
such as intestine and kidney in rainbow trout (Wright et al., 2004). Moreover, Paoactivity
has been studied in the liver, kidney, uterus, brain, lung, and skeletal muscles of rats
(Pavlov et al., 1991), humans (Suzuki et al., 1984) and catfish (Kumazawa and Suzuki,
1987). The ubiquity of key enzymes for polyamine metabolism supports the importance
of polyamines across tissue types and organisms. It is necessary to know if Arg, Odc and
Pao are present in the tissues and organs of killifish. Only limited information on the
tissue distribution is available at present; therefore, the influence of hypoosmotic
challenges on the enzymatic activities of Arg, Odc and Pao was investigated in multiple
tissues including gill and intestine (osmoregulatory organs), liver, heart and muscle
(energetic metabolism organs), spleen(immune system organ) and testes (reproductive
organ).
4.2 Materials and methods
4.2.1 Experimental animals
Adult F. grandis were collected from a local hatchery (Gulf Coast Minnows,
Thibodeaux, LA, USA) and maintained in a 330-L aquarium with a single recirculation
system containing mechanical and biological filtration and UV-sterilization. Salinity was
maintained at 5 ±< 0.1 ppt water, which was prepared by diluting Instant Ocean®
(Blacksburg, VA, USA) with reverse osmosis water. Temperature was kept at 23 ± 1˚C,
and salinity and temperature were monitored daily using a water quality meter (YSI,
73
Yellow Spring, OH, USA). Partial replacement of system water was performed at least
twice a week to maintain ammonia and nitrite below the detection levels of commercial
kits (Aquarium Pharmaceuticals, Chalfont, PA, USA). Dissolved oxygen was kept near
air-saturation using a low-pressure aerator. A 12 h light:12 h dark cycle was provided by
fluorescent lighting. Fish were fed a commercial pellet (Cargill, Franklin, LA, USA)
twice daily at 2% body weight per day throughout the study.
4.2.2 Experimental design
Sixty adult fish weighing between 7.8 and 11.5 g were acclimated to 5 ppt water
before being randomly divided into two groups. Following acclimation, fish were
transferred to aquaria containing either 5 ppt or 0.1 ppt water for one week and fed twice
daily until 12 h before sampling. Following exposure, fish were killed by caudal
severance and testes, gills, intestine, liver, heart, cranially-located epaxial muscle, and
spleen were collected from each fish. The intestinal tract from the end of the pyloric
sphincter to the anus was sampled as the whole intestine. Tissues (N=6 for each salinity)
were immediately stored in at least 10-times volume of RNAlater (Invitrogen, Carlsbad,
CA, USA), stored at 4 oC overnight, and transferred to a -20 oC freezer until total mRNA
extraction. Distribution of arg II and odc mRNA levels in tissues of adult F. grandis
acclimated to 5 ppt water was analyzed using RT-PCR. PCR products were run on a 2%
agarose gel to confirm the presence of a single product. All cDNA samples were
subsequently used in qPCR to assess the relative tissue distribution of arg II and odc
mRNA in tissues, as described in Section 4.2.3. Tissues were flash frozen in liquid
nitrogen and stored at -80 oC until the activities of arginase, Odc, and Pao were measured,
as described in Sections 4.2.4, 4.2.5, and 4.2.6, respectively.
74
4.2.3 arg II and Odc mRNA levels
TRIzol reagent (Invitrogen, Carlsbad, CA, USA) was used for the extraction of
total RNA from tissues according to the manufacturer’s instructions. Expressed sequence
tags (EST) for arg II (AB290198.1) and odc (CN977303.1) from F. heteroclitus were
obtained using the National Center for Biotechnology Information database
(http://www.ncbi.nlm.nih.gov/) and used to derive forward and reverse primers for
quantitative PCR (Integrated DNA Technologies Inc., Coralville, IA, USA). qPCR
analysis was performed for the target genes, arg II and odc, and 18S ribosomal RNA (18s
rRNA) was used as the reference gene. Analysis was conducted using a commercial kit
(SYBR Green PCR Core Reagent, Applied Biosystems, Carlsbad, CA, USA) with an
ABIPRISM 7000 Sequence Detection System (Carlsbad, CA, USA). Reagents included
5 µL cDNA (0.1 µg), 2 µL forward primers (10 nM), 2 µL reverse primers (10 nM), 5 µL
SYBR Green reagent, and 6 µL Ultrapure water (Invitrogen, Grand Island, NY, USA) in
a total volume of 20 µL. Samples were cycled 40 times at 95˚C for 15 s and 60˚C for 1
min. Relative changes in mRNA levels were made using the 2 -∆∆Ct method (Fleige et al.,
2006).
4.2.4 Total Arginine activity in multiple tissues
Total arginase activity was measured as described previously (Mommsen et al.,
1983). Briefly, tissues were homogenized in HEPES buffer (50 mM, pH=7.5) (V:V for
1:5) and centrifuged at 12,000 g for 4 min at 4 ˚C. Supernatants were incubated in 50
mM MnCl2 for 10 min, and the pre-incubated supernatant was mixed with reaction
reagent (250 mM L-arginine, 1 mM MnCl2, and 50 mM HEPES (pH=8.0). After 30 min,
25 µL 70% perchloric acid was added to the reaction mix and then neutralized with
75
KHCO3. The enzymatic reaction was performed at room temperature and the reaction
mix was centrifuged at 12,000 g for 4 min at 4 ˚C. The supernatant was analyzed for
total urea using a QuantiChrom urea assay kit (Bioassay System, Hayward, CA, USA),
and total arginase activity was expressed in µmol urea/h/ g tissue.
4.2.5 Decarboxylase activity in multiple tissues
The activity of ornithine decarboxylase did not differ in fresh tissue or in tissues
flash frozen at the time of collection in liquid nitrogen and stored at -80˚C data not
shown). Therefore, all Odc activity assays were performed on frozen gill samples, which
were thawed on ice and homogenized in reaction buffer (50 mM K2HPO4 buffer (pH 7.5),
5 mM dithiothreitol, 0.2 mM EDTA, 0.05 mM pyridoxal 5-phosphate). Samples were
centrifuged at 20,000 g for 20 min at 0 ˚C. The supernatant was added to a 16 mm x 100
mm borosilicate tube with an airtight seal formed with a double seal rubber stopper
(Kimble Chase Kontes, K-882310) and infiltrated by a plastic centerwell (Kimble Chase
Kontes, K-882320). The centerwell contained 1.3 cm2 of Whatman #1 filter paper
saturated with 133 µL 40% KOH. A 0.5 µCi mixture of DL-[1-14C] ornithine
hydrochloride (52 mCi/mmol, Moravek Biochemicals, Brea, CA, USA) and cold Lornithine (Sigma, St. Louis, MO, USA) were added anaerobically to the supernatant. The
ornithine concentration used in the assay mixture (1.2 mM) was the same as used in a
previous study (Watts et al., 1996). The reaction mixture was kept in a water bath at 25
˚C for 90 min, and 5% trichloroacetic acid was added to stop the reaction. Radiolabeled
14
CO2 was collected overnight on filter paper saturated with 40% KOH. The filter paper
was placed in 5 mL Ultima Gold cocktail (Perkin-Elmer, Waltham, MS, USA), and total
14
CO2 was measured using a Beckman LS 6500 Liquid Scintillation Counter (GMI Inc.
76
Ramsey, Minnesota, USA) as described by Watts (Watts et al., 1996). Internal
standardization demonstrated both constant and negligible 14C quench, so no quench
correction was applied to the data analysis. Odc activity was expressed as nmol
CO2 produced/h/g tissue.
4.2.6 Testing polyamine oxidase (Pao) activity in different tissues
Tissues were homogenized in a 15-fold volume of MilliQ water with
homogenizer (PRO 200, Lab depot Inc. Dawsonville, GA, USA). The crude homogenate
was used as a polyamine oxidase source. The assay method for Pao was modified from
the protocol description by Suzuki (Suzuki et al., 1984), in which Pao catalyzed
homovanillic acid to a highly fluorescent compound. The fluorescence intensity was
measured with excitation at 355 nm and emission at 460 nm using a VICTORTM X3
multi-label plate reader (Perkin Elmer, Santa Clara, CA, USA). The assay mixture
without substrate (N1-acetylspermine trihydrochloride) was incubated as a blank control,
and 2.2 nmol H2O2 was added instead of homovanillic acid as an internal standard.
4.2.7 Statistical analyses
One-way ANOVA and the Tukey post hoc test were performed using SAS (v. 9.3,
SAS Institute Inc., Cary, NC, USA) to test for significant differences within tissues in
both the 5 ppt and 0.1 ppt treatments. Statistical differences were considered significant
at P-values ≤0.05.
4.3 Results
4.3.1 Tissue distribution of arg II and odc
The mRNA levels of arg II and odc were measured in the gills, intestine, testes,
heart, muscle, liver, and spleen relative to 18s rRNA. All mRNA levels were normalized
77
to mRNA level of the gill in 5 ppt-acclimated fish. PCR primers used to amplify arg II
(Figure 4.1 panel A) and odc (Figure 4.2 panel A) produced single amplicons in all
tissues of adult male killifish, as demonstrated by 2% agarose gel electrophoresis of PCR
products. Quantitative PCR was subsequently used to assess the relative tissue
distribution of arg II (Figure 4.1 panel B) and odc (Figure 4.2 panel B) following 7 d
exposure of F. grandis to either 5 ppt or 0.1 ppt water. ArgII mRNA transcript was
highly expressed in the gills, regardless of salinity (P<0.05), compared to all other tissues
(Figure 4.1 panel B). Moreover, arg II mRNA levels increased significantly only in the
gills (P=0.0459), liver (P=0.0099) and spleen (P=0.0106) of fish acclimated to 0.1 ppt
water compared to the 5 ppt controls (Figure 4.1 panel B).
Although odc mRNA was most highly expressed in the testes of male killifish,
there were no significant differences in mRNA levels between the testes and gills in 5 ppt
or 0.1 ppt-acclimated fish (Figure 4.2 panel B). In contrast, odc mRNA levels were much
higher in the gills than in other tissues, except for the testes in both 5 ppt and 0.1 ppttreated fish. Low odc mRNA levels were observed in the spleen, muscle, and heart. Odc
mRNA levels were not significantly different from one another in all observed tissues
between 5 ppt and 0.1 ppt –acclimated fish at 7 d post-transfer (Figure 4.2 panel B).
4.3.2 Arginase (Arg) and ornithine decarboxylase activities in tissues of F. grandis
After determining the mRNA levels of arg II and odc genes, Arg and Odc
activities were analyzed. Despite the fact that the gills had the highest abundance of arg
II mRNA transcripts of all the tissues analyzed, the liver had a significantly higher level
of Arg activity (P<0.0001 for both salinities). The testes, intestine, muscle, and spleen
had a similar level of Arg activity (P>0.05 for both salinities), whereas the heart had a
78
A
bp
SM G T
I
L H
M
S
200
150
100
B
1.4
0.1 ppt
5 ppt
A*
Relative arg II mRNA level
1.2
a
b
B
1.0
0.8
0.6
*
0.4
0.2
0.0
*
e
n
rt
le
er
es
lls
Gi Test testin Liv Hea usc Splee
M
In
lls
Gi
t
r
s
e
e
ste
tin Live Hear uscl pleen
Te Intes
S
M
Figure 4.1: (A) Semi-quantitative assessment of arg II mRNA in the tissues of F. grandis
acclimated to 5 ppt water using RT-PCR analysis. SM represents 50 bp size marker, and
G, T, I, L, H, M, and S represent the gills, testes, intestine, liver, heart, muscle, and
spleen, respectively. (B) Relative arg II mRNA levels in tissues of F. grandis acclimated
to 5 ppt or 0.1 ppt water for 7 d (N = 6 per salinity). 18S ribosomal RNA was used as a
reference gene, and arg II mRNA levels were normalized to the mRNA level of the gills
(5ppt-acclimated fish) and assigned a value of 1. Lowercase letters indicate a significant
difference (P ≤ 0.05) in comparison to gills at 5 ppt for all tissues; uppercase letters
indicate a significant difference (P ≤ 0.05) in comparison to gills in 0.1 ppt exposed fish;
asterisks indicate a significant difference (P ≤ 0.05) in comparison to the 5 ppt control.
79
A
bp
SM G T
I
L H
M
S
200
150
100
B
1.8
0.1 ppt
5 ppt
Relative odc mRNA level
1.6
A
1.4
A
a
1.2
a
1.0
0.8
0.6
B
b
0.4
0.2
0.0
e
n
rt
ls
le
er
es
Gil Test testin Liv Hea usc Splee
M
In
e
n
rt
ls
le
er
es
Gil Test testin Liv Hea usc Splee
M
In
Figure 4.2: (A) Semi-quantitative assessment of odc mRNA in tissues of F. grandis
acclimated to 5 ppt water using RT-PCR analysis. SM represents 50 bp size marker; G, T,
I, L, H, M, and S represent the gills, testes, intestine, liver, heart, muscle and spleen,
respectively. (B) Relative odc mRNA levels in tissues of F. grandis acclimated to 5 ppt
or 0.1 ppt water for 7 d (N = 6 per salinity). 18S ribosomal RNA was used as a reference
gene, and odc mRNA levels were normalized to the mRNA level of the gill (5pptacclimated fish) and assigned a value of 1. Lowercase letters indicate a significant
difference (P ≤ 0.05) in comparison to the gill at 5 ppt for all tissues; uppercase letters
indicate a significant difference (P ≤ 0.05) in comparison to the gill in 0.1 ppt water
exposed fish; asterisks indicate a significant difference (P ≤ 0.05) in comparison to the 5
ppt control.
80
significantly lower level of Arg activity (P<0.0001 for both salinities) compared to the
gills (Figure 4.3). Hypoosmotic exposure only increased Arg activity in the gills
(P=0.0184) compared to the 5 ppt control, but did not affect enzymatic activity in other
Arginase activity (mol urea produced/ h / g tissue)
tissues (Figure 4.3).
b
B
600
400
A*
a
a
a
A
a
a
200
0
lls
Gi
A
A
C
c
t
r
s
e
e
ste
tin Live Hear uscl pleen
Te Intes
S
M
lls
Gi
A
t
r
s
e
e
ste
tin Live Hear uscl pleen
Te Intes
S
M
Figure 4.3: Arginase activity in the tissues of adult F. grandis acclimated to 5 ppt and 0.1
ppt water for 7 d. Data are represented as mean values ±SEM (N = 6) in µmol urea
produced /h/g tissue. Lowercase letters indicate a significant difference in levels of
different tissues (P ≤ 0.05) in comparison to the level of gills in 5 ppt treated fish;
uppercase letters indicate a significant difference in levels of different tissues (P ≤ 0.05)
in comparison to the gill in 0.1 ppt treated fish; asterisks indicate a significant difference
(P ≤ 0.05) in the tissues from the 0.1 ppt exposed fish in comparison to those from the 5
ppt-acclimated fish.
There was no significant difference in Odc activity between all tissues measured
in 5 ppt-acclimated fish (Figure 4.4). After transfer to 0.1 ppt water, the gills had higher
Odc activity than all other tissues except the testes (Figure 4.4). Hypoosmotic transfer
increased Odc activity in the gills (P=0.0044), but did not affect Odc activity in other
tissues (Figure 4.4).
81
Odc activity (nmol CO2 produced / h / g tissue)
1.0
A
.8
A*
.6
B
a
a
.4
.2
0.0
Gi
lls
t
r
s
e
e
ste
tin Live Hear uscl pleen
Te Intes
S
M
e
n
rt
le
er
es
lls
Gi Test testin Liv Hea usc Splee
M
In
Figure 4.4: Ornithine decarboxylase activity in tissues of F. grandis acclimated to 5 ppt
and 0.1 ppt water for 7 d. Data are represented as mean values ±SEM (N = 6) and
expressed as nmol CO2 produced/h/g tissue. Lowercase letters indicate a significant
difference in levels of different tissues (P ≤ 0.05) in comparison to the level of gill in 5
ppt treated fish; uppercase letters indicate a significant difference in levels of different
tissues (P ≤ 0.05) in comparison to the gill in 0.1 ppt treated fish; asterisks indicate a
significant difference (P ≤ 0.05) in the tissues from the 0.1 ppt exposed fish in
comparison to those from the 5 ppt-acclimated fish.
4.3.3 Pao activity in multiple tissues
To assess the extent of polyamine catabolism, Pao activity in multiple tissues of
F.grandis acclimated to 5 ppt and 0.1 ppt water was measured. After 7 d acclimation to 5
ppt and 0.1 ppt water, the intestine, liver, and gills had higher Pao activity than that in
other tissues, as shown in Figure 4.5. Hypoosmotic exposure increased Pao activity in
the gills (P=0.0094), but did not affect the Pao activity of other tissues, compared to
tissues from 5 ppt-acclimated fish (Figure 4.5). The gill had the highest Pao activity
compared to all other tissues during hypoosmotic exposure.
82
Pao activity (nmol / g tissue / 30 min)
400
A*
A
300
a
a
C
200
c
100
B
b
0
b
b
B
b
e
n
rt
ls
le
er
es
Gil Test testin Liv Hea usc Splee
M
n
I
B
B
e
n
rt
ls
le
er
es
Gil Test testin Liv Hea usc Splee
M
n
I
Figure 4.5: Pao activity in tissues of F. grandis after 7 d of hypoosmotic exposure. Data
are represented by mean ±SEM (N=6) and expressed as nmol/g tissue /30 min.
Lowercase letters indicate a significant difference in levels of different tissues (P ≤ 0.05)
in comparison to the level of gill in 5 ppt treated fish; uppercase letters indicate a
significant difference in levels of different tissues (P ≤ 0.05) in comparison to the level of
gill in 0.1 ppt treated fish; asterisks indicate a significant difference (P ≤ 0.05) in the
tissues from the 0.1 ppt water exposed fish in comparison to those from the 5 pptacclimated fish.
4.4. Discussion
4.4.1 The enzymatic activity of arginase, ornithine decarboxylase and Pao in the gills
This study suggests that changes in enzymatic activity associated with polyamine
metabolism in gills may be an important response of euryhaline fish to acute
hypoosmotic exposure. The fish gill is a multipurpose organ that plays dominant roles in
osmotic and ionic regulation, acid-base regulation, gas exchange, and excretion of
nitrogenous wastes (Evans et al., 2005; Wood and LeMoigne, 1991; Wood and Marshall,
1994). In F.grandis, hypoosmotic challenges increased the enzymatic activities of Arg,
Odc and Pao only in the gills (Figures 4.3, 4.4 and 4.5). This indicates that although all
83
tissues had measureable levels of enzymatic activities associated with polyamine
metabolism, only the gill had enzyme activities affected by hypoosmotic challenge. The
intracellular concentrations of putrescine, spermidine, and spermine are strictly regulated
by multiple biochemical pathways involving Arg, Odc, and Pao, suggesting polyamines
play an important role in various of physiological functions in plants and animals
(Galston, 1983; O'Brien et al., 1975; Sekowska et al., 1998). Putrescine participates in
modulation of gene transcription, signal transduction, membrane stability and receptorligand interactions; spermidine participates in modulation of gene translation and
chromatin structure; and spermine is involved in modulation of mRNA/DNA stabilization,
cell growth and proliferation (Pegg and McCann, 1982; Ray et al. 1999; Tabor and Tabor,
1984). This study suggests that hypoosmotic exposure stimulates polyamine biosynthetic
pathways, resulting in an increase in putrescine, spermidine and spermine concentrations
in the gill (Chapter 2). Highly cationic polyamines such as spermidine and spermine
mediate early phase compensatory responses in the euryhaline killifish when faced with
osmotic challenges. Although gill spermidine and spermine increased transiently in
killifish with freshwater exposure, putrescine remained elevated over time, suggesting a
possible role of this polyamine in acclimation of killifish to fresh water. Moreover,
spermidine, and spermine concentrations increased in the gills of Callinectes danae
during during hyperosmotic exposure (Silva et al., 2008). Elevated putrescine and
spermidine concentrations were also present in the gills of Callinectes sapidus after
hyperosmotic exposure (Lovett and Watts, 1995). In contrast, the putrescine
concentration of Artemia nauplii decreased with an increase of environmental salinity
(Watts and Good, 1994). Although there was no effect on spermidine and spermine
84
concentration, Odc activity and putrescine concentration increased in brine shrimp during
hypoosmotic exposure (Watts et al., 1996). Persumably, salinity challenges stimulated
perturbation of polyamine homeostasis in order to mediate a series of biological functions,
such as stimulating cell apoptosis (Laurent et al., 2006). In this study, hypoosmotic
exposure resulted in upregulation of key enzymes of polyamine metabolism, although
additional studies are needed to determine the physiological effects of individual
polyaminesto hypoosmotic exposure. Surprisingly, the increase in Odc activity during
hypoosmotic exposure was not the result of an increase in odc mRNA level at 7 d
hypoosmotic exposure (Figures 4.2 and 4.4). Actually, Odc activity is regulated by
translational as well as post-translational mechanisms (Lovkvist et al., 2001). This
phenomenon can be explained mainly by post-transcriptional regulation. Previous
studies have also shown that hypoosmotic challenge increased Odc activity without any
detectable change in Odc mRNA level in L-1210 cell line (Poulin and Pegg, 1989). As
such, the induction of Odc activity can be explained partly by an increased efficiency of
Odc mRNA translation.
4.4.2 The enzymatic activity of arginase, ornithine decarboxylase and Pao in testes,
intestine, liver, heart, muscle and spleen
Hypoosmotic exposure did not affect the activities of Arg, Odc and Pao in testes,
intestine, liver, heart, muscle and spleen. Among those tissues, liver had the highest
arginase activity in F.grandis (Figure 4.3) in both salinities, in accordance with earlier
results of arginase activity in other fish species (Cvancara, 1969). However, the liver had
a low expression of arg II mRNA (Figure 4.1 panel B). Morris (2004) suggests that Arg
I, not Arg II, is the primary arginase protein in the liver. In addition, studies have shown
that arg I mRNA levels are higher than the levels of arg II in the liver of adult rainbow
85
trout (Wright et al., 2004), humans (Morris et al., 1997), and Xenopus tadpoles (Patterton
and Shi, 1994). Thus, Arg I mainly contributed to liver arginase activity. There is no
evidence of Arg I in Fundulus, and most teleost fish are obligate ammonioteles lacking a
functional OUC beyond embryogenesis; thus, Arg II is the major functional arginase in
them to produce ornithine (Wright et al., 2004; Wu et al.,1998). Moreover, as a
substreate of arginase, arginine is a precursor of polyamines and various amino acids,
such as proline and glutamate. Arginase activity in fish is found in high abundance in
tissues with high metabolic activity, such as the liver, because these tissues play an
important role in amino acid metabolism in fish (Ketola, 1982).
Another key enzyme for polyamine biosynthesis is Odc. In this study, Odc activity
was higher in the testes compared to all other tissues, excluding the gills in 0.1-ppt
acclimated killifish (Figure 4.4). Odc is the rate-limiting enzyme for polyamine
biosynthesis, and polyamines, namely putrescine, spermidine and spermine, are present in
and synthesized by germ cells, Sertoli, and Leydig cells of the testis in the rat (Lefèvre et
al., 2011). Moreover, this study shows that the liver and intestine had high Pao activity
in F. grandis tissues (Figure 4.5). The intestine had higher Pao activity for both salinities
at 7 d, which is consistent with the high Pao activity observed in the intestine of catfish
(Kumazawa and Suzuki, 1987). The intestine is responsible for water and food
absorption, and this action can compensate for water loss from kidney (Evans et al.,
2005). Thus, intestine is an important organ for osmotic regulation. Besides the intestine,
liver has high Pao activity because it functions in lipid storage and works as an enzymatic
pool in other species (de la Torre et al., 1999).
86
In summary, biosynthetic and catabolic pathways tightly regulate polyamine
levels by multiple tissues. All polyamines components played a role in dealing with
hypoosmotic challenge; however, spermidine and spermine levels dropped after 1 d. Gill
putrescine stored at early and later compensatory response to hypoosmotic challenge.
High Arg, Odc and Pao activity in the gills, intestine and liver are presumably involved in
polyamine homeostasis in the killifish. The ratio of putrescine level over the sum
concentration of spermidine and spermine reached its maximum at 7 d post-transfer,
suggesting more putrescine accumulated in the gill at later compensatory response to
hypoosmotic challenge (Chapter 2). This study indicates putrescine may play a role in
the recovery F.grandis from hypoosmotic exposure. Gill remodelling is a strategy for
killifish survival in hypoosmotic exposure, although a link between putrescine and gill
remodelling needs clarification. Therefore, the role of putrescine in gill remodelling
during hypoosmotic exposure should be further studied.
87
CHAPTER 5: CONCLUSION
Polyamines are a family of low molecular weight organic cations, and they have
been implicated in diverse physiological processes. My research to date has described a
possible role of polyamines in the acute response of the gulf killifish, Fundulus grandis,
to hypoosmotic exposure. The purpose of this study was to describe the transcription and
enzymatic activities of key enzymes for polyamine metabolism, to assess polyamine
levels during this process, and to assess the putative roles of polyamines in the gills of F.
grandis during hypoosmotic challenges. Also, the influence of the irreversible inhibitor
ornithine decarboxylase (Odc) by alpha-DL-difluoromethylornithine (DFMO) was used
to gain insight to the influence of impairment of polyamine biosynthesis on the
physiological response of killfish to freshwater exposure. The current study sought to
answer the following questions: a) Do osmotic challenges influence the synthetic and
catabolic pathways that regulate gill polyamine concentrations? b) Do polyamines play a
role in gill epithelial remodeling during hypoosmotic exposure? c) Does polyamine
accumulation play a role in Cl- secretion associated with cell swelling in vitro during
hypoosmotic exposure? and d) Does polyamine metabolism occur in multiple tissues
during hypoosmotic challenges in killifish?
Major empirical findings from my first study showed that a large increase in
mRNA levels and the enzymatic activities of Arg and Odc in the gills occurred in
response to exposure of killifish to low environmental salinity (Figures 2.1 and 2.2). One
of the most significant results to emerge from this study was that odc mRNA levels
increased exponentially as environmental salinity decreased at all observed time points
(data not shown). Moreover, the levels of spermidine and spermine were significantly
88
elevated in the gills at only 1 d exposure to 0.1 ppt, whereas the putrescine level was
significantly elevated by this treatment at 1 d and beyond (Figure 2.3). The ratio of
putrescine level over the sum of spermidine and spermine levels increased progressively
following transfer to 0.1 ppt water (Table 2.2). This change in this ratio is associated
with a progressive reduction in the concentrations of spermidine and spermine, yet a
sustained elevation in the concentration of putrescine. This change in the ratio also
suggests that both spermine and spermidine mediated the acute compensatory response of
hypoosmotic challenges in the killifish; whereas, putrescine play an important role in the
prolonged compensatory response of hypoosmotic shock in the euryhaline gulf killifish.
Gill putrescine reached its maximum concentration at 7 d post-transfer to hypoosmotic
environment, concommitant with an increase in polyamine oxidase activity (Figure 4.5),
suggesting that polyamine catabolism was upregulated following hypoosmotic exposure
(0.1 ppt). The polyamine catabolism was not measured at early time points; therefore, it
cannot conclude how polyamine catabolism changed. However, polyamine levels should
be kept in a homeostasis state, and polyamine catabolism has a high possibility to
upregulated initially. Polyamine catabolism is a back-conversion pathway, converting
highly aminated polyamines, such as spermine and spermidine, back to putrescine. These
results showed that osmotic challenges affected gill polyamine levels by regulating both
polyamine biosynthesis and catabolism. Putrescine, spermidine and spermine may play a
role in different temporal phases after freshwater exposure.
This study showed that the efficacy of DFMO on Odc activity was dependent on
exposure salinity. Odc activity and polyamine concentrations were significantly reduced
by DFMO after 0.1 ppt transfer; however, DFMO had no effect on endogenous Odc
89
activity in fish gills in the absence of hypoosmotic exposure (Table 2.3; Figures 2.4 and
2.5). Surprisingly, gills still exhibited a marked reduction in gill polyamine
concentrations with DFMO despite the inability to inhibit the enzyme. This result may
be explained by the polyamine-mediated feedback of Odc activity. Polyamines exerted
an inhibitory effect on Odc synthesis, and they also stimulated Odc degradation by
inducing antizyme synthesis. Another explaination may be the existence of DFMOinsensitive Odc. A large component of total Odc activity in the killifish gill is DFMOinsensitive Odc, especially the endogenous sources not stimulated by hypoosmotic
exposure.
Another significant finding to emerge from this study was a strong correlation
between the increase of arg II and odc mRNA levels and the morphological transition in
MRCs during hypoosmotic exposure (Figures 3.2 and 3.3). This strongly suggests that
upregulation in polyamine biosynthesis may either mediate some of the phenotypic
changes to the gill epithelium or that it is a consequence of the cellular swelling and/or
damage during the early phases of freshwater transfer. Hypoosmotic challenges induce a
morphological transition from seawater-type to freshwater-type MRCs in the gill
epithelium (Figure 3.2). Concurrent with the onset of morphological changes to the gill,
FW exposure elicited increases in the mRNA levels of c-fos, c-myc oncogenes, and
caspase-3 activity (Figure 3.7). These results suggested that polyamine-mediated cell
proliferation and apoptosis may be involved in gill remodeling after freshwater exposure.
Moreover, DFMO application reduced not only caspase-3 activity, but also the levels of
gill putrescine, spermidine and spermine after hypoosmotic exposure, indicating a
relationship between polyamines and cell apoptosis in the low environmental salinity.
90
The next goal of this thesis was to assess a possible association between gill
polyamine accumulation and cell volume disturbances or regulated recovery from cell
volume disturbance. This work was conducted by supplementing isolated opercular
epithelia exposed to hypotonic conditions with putrescine, spermidine, or spermine,
either singly or as a polyamine mixture. Opercular epithelium was used for this study
because it shares many similar properties to the gills, including the presence of MRCs,
the existence of a transcellular Cl- secretory mechanism similar to that found in MRCs of
marine fish gill, and has the capacity to undergo regulatory volume decrease in the face
of hypotonic swelling. As a proxy of cell swelling, short circuit current (Isc) was used to
reflect Cl- secretion across the epithelium. A hypotonic shock was applied on the
basolateral side of opercular epithelia. A decrease of 60 mOsM/Kg reduced Isc by
approximately 60% and hypotonic inhibition of Isc was accompanied by reduction in
membrane conductance (Gt). Administration of spermidine to the basolateral side of the
opercular epithelium prevented a decrease in Gt and Is across the epithelium induced by
hypotonic exposure (Figures 3.4, 3.5 and 3.6). Thus, polyamine accumulation may
aggravate the cell swelling by inhibiting Cl- secretion in vitro after hypotonic exposure.
Finally, this study shows that the transcriptional and enzymatic activities of Arg, Odc and
Pao were upregulated in the gills, intestine and liver during acute freshwater exposure,
suggesting that polyamine levels are regulated on the whole-animal level in killifish
(Figures 4.1, 4.2, 4.3, 4.4,and 4.5). Intestine is another important osmoregulatory organ,
and it gains extra water through food ingestion to compensate for renal elimination after
transfer to fresh water. Low activities for Arg and Odc, and a high activity of Pao
suggest the intestine plays an important role in polyamine metabolism. Pao participates
91
in the regulation of putrescine concentration during extended freshwater exposure,
supporting a possible role of putrescine in maintenance of the FW gill phenotype.
These studies have offered a comprehensive perspective on the regulation of
polyamine metabolism in killifish. Increases in putrescine, spermidine and spermine
during osmotic challenges were observed in the killifish, and this pattern facilitated fish
to acclimate to fresh water by gill remodeling. Moreover, long-term exposure of killifish
to FW leads to the persistent elevation of gill putrescine concentration, but not in the
more highly aminated spermidine and spermine. This result suggested a relationship
between putrescine concentration in the gill and gill remodeling in the prolonged
compensatory response to hypoosmotic challenge. However, polyamine accumulation
may aggreavate cell swelling in MRCs by preventing Cl- secretion in the face of
hypotonic shock, which may be related to the functions of Na+/K+/2Cl- co-transporter
(NKCC) and cystic fibrosis transmembrane conductance regulator (CFTR) during
hypotonic shock (Hoffmann, 1992; Hoffmann, 2000), as previous studies have shown.
Therefore, it is premature to conclude whether polyamines are protective or cytotoxic to
cells during hypoosmotic challenges, although the regulation aimed at maintenance of gill
putrescene during FW acclimation supports an important role of this molecule.
Future studies should establish a direct linkage between arg II and odc mRNA
expression and polyamine concentration in the gill. Second, a direct relationship between
gill polyamine levels and cell apoptosis should be estabalished. Third, although the study
showed a correlation between polyamine and c-fos and c-myc transcription, a causal link
between them needs to be clarified during hypoosmoitc exposure. Forth, DFMO
application prevented a decrease in plasma Na+ and Cl- concentrations in the gill after
92
hypoosmoitc challenge, but clearly understand the the causal relationship between
polyamines and plasma osmolality needs further work.
93
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APPENDIX: PERMISSION OF REPRINT
Dear Ying Guan,
Thank you for your recent email requesting permission to reuse all or part of your
article in a new publication, a thesis or as part of your teaching.
The information first appeared as Guan, Y., Meng, Y.L. and Galvez, F. (2012).
The effects of salinity on gene expression of ion transporters in gulf killifish (Fundulus
grandis). International proceedings of chemical, biological& environmental engineering.
49:166-177. Reprinted by permission.
Please see permissions information at our website, and let us know if you have further
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VITA
Ying Guan was born in Heilongjiang, China, in 1980, the daughter of Shitang
Guan and Fengyun Li. After completing her degree at Boli High School, Heilongjiang,
in 1995, she entered the Qiqihaer Medical University, receiving the degree of Bachelor of
Clinic Medicine, 2003. She entered the Graduate School in the Department of Biological
Sciences at the Louisiana State University in August, 2007, receiving a Master's of
Statistics in May 2012.
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