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First published online August 17, 2007
Journal of Experimental Biology 210, 3126-3132 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.004150
Functional identification of an osmotic response element (ORE) in the promoter region of the killifish deiodinase 2 gene (FhDio2)
Departamento de Neurobiologia Celular y Molecular, Instituto de Neurobiología, Universidad Nacional Autónoma de México, Quéretaro, Qro. CP 76230, México
* Author for correspondence (e-mail: lucianikolaia{at}gmail.com)
Accepted 5 June 2007
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Summary |
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Key words: osmotic response element, osmotic stress, deiodinase type 2 gene, Fundulus heteroclitus, thyroid hormones
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Introduction |
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). The first
osmotic response element (ORE) in a eukaryotic genome was originally described
in the rabbit aldolase reductase gene
(Ferraris et al., 1996),
followed by the characterization of its corresponding transcription factor,
the ORE binding protein (OREBP) (Ferraris
et al., 1999). Tyrosine phosphorylation is required for OREBP to
translocate into the nucleus and activate transcription. Although the pathway
responsible for OREBP phosphorylation remains to be fully elucidated, tyrosine
kinases p38 and protein kinase A (PKA) have been implicated
(Ferraris et al., 2002). In
marked contrast to terrestrial vertebrates, information regarding osmotic
stress responses in aquatic species is scarce and fragmented. Up to now OREBPs
have been amply assessed during hyperosmotic conditions, but these studies
have been mainly restricted to in vitro experiments and to mammals.
Recently the transcriptional regulation in response to changes in
environmental salinity has begun to be explored in euryhaline fishes. Two
osmotic stress transcription factors have been identified in tilapia gill, the
osmotic transcription factor 1 (Ostf1) and the tilapia homolog of
transcription factor II B (TFIIB). Both show a specific and transient
up-regulation during hyperosmotic stress
(Fiol and Kultz, 2005). Ostf1
has strong homology to the mammalian glucocorticoid leucine zipper protein;
however, no steroid is required for its induction during hyperosmotic,
NaCl-mediated stress in fish (Fiol et al.,
2006).
Because of their binding to nuclear receptors, thyroid hormones (TH) are
well-known transcription mediators. TH play a central role in regulating
diverse homeostatic functions, from the basal metabolic rate and protein
synthesis, to development and cell differentiation
(Anderson et al., 2000).
Nevertheless, the function of TH in hydro-osmotic balance in fish has been a
controversial issue (for reviews, see
McCormick, 2001;
Orozco and Valverde-R., 2005;
Klaren et al., 2007). Previous
data from our laboratory have shown that in the euryhaline teleost
Fundulus heteroclitus (Fh), hypo-osmotic stress elicits an increase
in liver iodothyronine deiodinase type 2 (D2) activity
(Orozco et al., 1998). D2 is
known to be the major provider of T3 at the target cell level;
thus, an up-regulation of this enzyme after an osmotic challenge suggested the
participation of TH during this homeostatic response. In the present study we
used a bioinformatic approach (Heinemeyer
et al., 1998) to identify two conserved ORE motifs within the
5' UTR of the FhDio2 gene
(Orozco et al., 2002a). We
then carried out a series of in vivo and in vitro
experiments to examine the possible participation of these ORE motifs in
regulating FhDio2 gene expression during hypo-osmotic stress in
Fundulus heteroclitus. Our data demonstrate the participation of a
putative OREBP-activated pathway in the liver of killifish challenged by
abrupt low salinity.
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Materials and methods |
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) with
the available vertebrate matrices and a threshold score up to 85.
Animals
Seawater-adapted male Fundulus heteroclitus L. (killifish), body
mass 4–6 g, were collected from the estuarine creeks of the Matanzas
River (St Augustine, FL, USA). After capture, fish were deparasitized and kept
in tanks with running seawater (SW) piped directly from the ocean at a
temperature of around 28°C. Animals were fed ad libitum (Silver
Cup, Nelson and Sons, Murray, UT, USA) and maintained on a light:dark cycle of
14 h:10 h. All experimental protocols used in this study were approved by the
Institutional Animals Ethics Committee. Experiments were carried out 1 week
after acclimatization.
In vivo experiments
For each experiment, fish (N=10–12) were placed into tanks
with running SW and constant aeration. At the start of the experiment, the
water supply was shut down, and the water volume of each tank was adjusted to
6 l with a 1:1 ratio of salt- and freshwater (50%). Except for the change in
water salinity, control and experimental groups were handled in the same
manner. Based on previous experiments
(Orozco et al., 1998), fish
were sacrificed 0.5, 1, 2, 4, 6, 8, 12 and 16 h after the hypo-osmotic
challenge. Animals were sacrificed by decapitation, and the liver was
collected and divided to measure D2 mRNA concentrations, D2 activity and for
nuclear extraction (see below). D2 mRNA was measured in pools.
In vitro experiments
Seawater acclimated fishes (N=2–3) were sacrificed, and
their livers were dissected and cut into small pieces of about 2 mm (5 mg wet
mass) in the presence of L-15 medium (Invitrogen, Carlsbad, CA, USA). Liver
explants were pooled, kept on ice for 30 min, and then randomly assigned to
either control or experimental 24-well plates. The L-15 medium in the
experimental plates was made hypotonic by simple dilution 1:1, restoring its
nutrients with an amino acid stock solution (Sigma-Aldrich, St Louis, MO, USA)
and galactose 10 g l–1. The explants were incubated for 6 h
in a metabolic bath chamber at 28°C under continuous agitation and
saturating humidity with 99% O2 and 1% CO2
(Janssens and Grigg, 1994).
Explants and the corresponding culture media were separately collected after
2, 4 and 6 h of incubation. Liver fragments were divided into two pools: one
to quantitate D2 activity and the other for nuclear extraction. D2 activity
was measured in tissue homogenates, and to evaluate tissue viability lactate
dehydrogenase (LDH) was measured in the culture media. For the tyrosine kinase
inhibitor experiments, explants were pre-treated with genistein 2 h prior
before exposure to L-15 hypo-osmotic medium.
Nuclear and cytoplasmic extracts
Control or experimental livers were resuspended in hypotonic buffer (10
mmol l–1 Hepes, pH 7.9, 10 mmol l–1 KCl, 1
mmol l–1 EDTA, 5 mmol l–1 DTT) as previously
described (López-Bojórquez
et al., 2004). The tissue was broken mechanically with a plastic
homogenizer at low velocity. To allow cell debris to settle the homogenates
were incubated at 4°C for 30 min, and the supernatants were collected. The
integrity of the nuclei was evaluated by Trypan-blue stain (1:1).
Nuclei were centrifuged at 800 g to separate them from the cytoplasmic fraction. The supernatants (cytoplasm) were diluted in an equal volume of dilution buffer (20 mmol l–1 Hepes, pH 7.9, 50 mmol l–1 KCl, 20% glycerol, 0.2 mmol l–1 EDTA, 0.5 mmol l–1 PMSF, 1 mmol l–1 DTT) for protein quantification. The pellet (nuclei) was recovered, resuspended in hypertonic buffer (50 mmol l–1 Tris-HCl, pH 7.9, 400 mmol l–1 NaCl, 400 mmol l–1 KCl, 10%, glycerol, 1 mmol l–1 EDTA, 5 mmol l–1 DTT, 0.5 mmol l–1 PMSF) and maintained with agitation at 4°C for 30 min. After centrifugation at 16 000 g for 25 min, the supernatant was resuspended in an equal volume of dilution buffer. Protein was quantified by the Bradford method (BioRad, Hercules, CA, USA). The same protocol was followed to isolate the nuclear and cytoplasmic fraction from the explants in the in vitro experiments.
Analytical procedures
LDH assay
The viability of liver explants was assessed by the quantitative
determination of LDH in the culture media using the reagents supplied in the
SPINREACT kit (SpinReact, Girona, Spain). The activity was normalized to the
protein content of the corresponding liver explants for each well. These data
were used to normalize D2 activity of the corresponding explant by multiplying
D2 specific activity and LDH activity. Normalized D2 results are expressed in
arbitrary units.
Deiodination assay
Enzyme activity was measured in duplicate as previously described
(Orozco et al., 2000).
Briefly, the total volume of the reaction mixture was 100 µl and contained
1 nmol l–1 125I-T4 [specific activity
1200 µCi µg–1 (1 µCi=3.7x1010 Bq);
NEN-Perkin Elmer, Wellesley, MA, USA], 25 mmol l–1 DTT
(Calbiochem, Darmstadt, Germany) and 100 µg/tube of liver homogenate
protein, and the assays were incubated for 1 h at 37°C. The released
acid-soluble 125I was isolated by chromatography on Dowex 50W-X2
columns (BioRad). Enzyme specific activity (fmol 125I
mg–1 h–1) was calculated as previously
described (Pazos-Moura et al.,
1991). Protein content was measured by the Bradford method. Since
the LDH activity in the culture medium is inversely proportional to cell
viability, the final D2 activity is reported as product of both enzymes, in
arbitrary units.
Measurement of D2 mRNA
For quantitation, RNA was isolated from livers (TRIzol, Invitrogen), and
cDNA was reverse-transcribed (Superscript, Invitrogen) from 10 µg
RNA using a gene-specific primer (TTCAGAGCTCATCTACTATCGT). The D2 mRNA
concentration was measured in duplicate by a competitive PCR as previously
described (García-G. et al.,
2004). The standard curve ranged from 104 to
109 molecules µl–1. The oligonucleotides used
(sense: CAAACAGGTGAAACTTGGCT and antisense: TCGTCGATGTAGACCAGC) amplified a
product of 270 bp (40 s at 94°, 40 s at 65°, 30 s at 72° for 35
cycles). Identical PCRs from the RNA samples prior to the reverse
transcription reaction yielded no detectable products, which indicates that
the RNA was not contaminated with genomic DNA. Results are expressed as
molecules µg–1 total mRNA used in the reverse
transcription.
Electrophoretic mobility shift assay (EMSA)
Putative OREBP-binding activity was studied by the use of
[32P]-labeled double-stranded oligonucleotides corresponding to
ORE1 (5' TGTAGA GGGAAAAGCT GGGACCA 3') or ORE2 (5'
TTTCAGG CGGAAAAGTA ACATTTC 3') (see below). The probes (200 ng in
1 µl) were labeled with 0.5 µl (5 U) of T4 phosphonucleotide kinase; 2.5
µl of buffer provided in the kit (Roche, Basel, Switzerland) and 3 µl of
32P-
ATP (NEN-Perkin Elmer, specific activity 3000 Ci mmol
l–1), and water to a final volume of 25 µl. The labeling
reaction was mixed with 1 ml of hybridization solution and then passed through
a nitrocellulose filter to eliminate excess 32P-ATP. The
quality of the labeled products was assessed by electrophoresis in denaturing
polyacrylamide gels exposed to photographic film. The binding reaction was
performed by incubating 10 µg of either nuclear or cytoplasmic protein with
the corresponding probes. The reaction mixture was incubated on ice for 40 min
in a buffer containing 20 mmol l–1 Hepes, 50 mmol
l–1 KCl, 20% glycerol, 0.2 mmol l–1 EDTA,
0.5 mmol l–1 PMSF, 1 mmol l–1 DTT, 1 µg
µl–1 BSA, and polydI/dC (Pharmacia, NY, NY, USA). The
reaction mixture was loaded onto a 5 % non-denaturing polyacrylamide gel and
resolved at 120 V for 4 h. The gel was dried, and the DNA–protein
complexes were visualized by exposure in a Storage Phosphor Screen (Molecular
Dynamics, San Francisco CA, USA). The screens were read in a
Phosphorimager (Storm Molecular Dynamics) and quantified with the
ImageQuant software (Molecular Dynamics). The radioactivity of each band was
corrected for the background and is presented in arbitrary units.
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In the D2 activity experiments, differences among groups were analyzed by one-way ANOVA coupled with a Bonferroni's multiple comparison test (control vs treatment). Differences were considered significant when P<0.05.
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), and ORE2 only differed in one nucleotide
(Fig. 1B).
Hypo-osmotic stress in vivo triggers the nuclear recruitment of a putative OREBP and up-regulates mRNA expression and D2 activity
To examine whether a putative OREBP is associated with FhDio2
transcription via the ORE elements identified in the 5'
promoter region of the gene, we performed EMSA with specific oligonucleotides
containing the endogenous ORE1 and ORE2 motifs identified above
(Fig. 1). Nuclear extracts from
both control and hypo-osmotically stressed fish were obtained at 0.5, 1, 2, 3,
4, 8, 12 and 16 h after the stress. As shown in
Fig. 2A, hypo-osmotic stress
was accompanied by clear-cut recruitment of cytoplasmic protein into the
nuclei. Indeed, this protein–DNA binding occurred in a biphasic mode: an
initial protein translocation 2 h after stress, and a second, more intense
wave of recruitment 6 h later. Furthermore, the nuclear protein–DNA
binding correlated with the parallel disappearance of protein from the
cytoplasmic compartment. The protein reappeared in the cytoplasm 12 h after
initiation of the experiment. Fig.
3 shows the specificity of the putative OREBP complex. Formation
of the complex was specifically blocked by competition with a 100-fold excess
of the identical but unlabeled oligonucleotide, as well as when nuclear
extracts were omitted from the binding reaction or when a random
oligonucleotide was used. Closely coupled in time to this initial putative
OREBP nuclear recruitment, both the D2 mRNA concentration and D2 activity
(Fig. 4) increased
significantly. D2 mRNA attained maximum values 8 h after stress, which
coincides with the second and more intense wave of the putative OREBP
recruitment. The increase in enzyme activity was slower and became significant
12 h after the osmotic challenge.
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Liver cells in vitro directly detect extracellular osmotic changes
Consistent with results from the in vivo experiments, when liver
explants were confronted directly with the hypo-osmotic challenge, there was a
significant protein translocation into the nuclei
(Fig. 5A). This
protein–DNA recruitment is first observed at 2 h and reaches a peak 4 h
post-stress. Furthermore, this conspicuous, putative OREBP-translocation
response is followed by a significant increase in hepatic D2 activity 2 h
later (Fig. 5B).
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) showing
that killifish hepatic D2 increases during hypo-osmotic stress. Indeed, the
present studies demonstrate that to cope with a sudden drop in salinity, the
liver of killifish triggers a homeostatic response in which a putative OREBP
kinase-activated pathway stimulates D2 transcription and enzymatic activity.
Furthermore, the participation of this putative signaling pathway is strongly
supported by the complete blockade of the response by a tyrosine kinase
inhibitor.
In mammals it is well known that hyperosmotic stress activates diverse
transcription factors, which in turn regulate key genes that maintain
hydrosmotic homeostasis (Ferraris et al.,
1999; Gatsios et al.,
1998). However, in teleosts, less is known about this
transcriptional regulation (Fiess et al.,
2007). Moreover, despite the fact that euryhaline fish are
constantly coping with osmoregulatory demands, available information about
hypo-osmotic stress is even scarcer. Our results clearly demonstrate that in
response to hypo-osmotic stress, at least one nuclear protein is recruited
into the ORE sequence present in FhDio2. Accordingly, we call this
protein a `putative OREBP'. Studies regarding OREBP activation under
hypotonicity have been scanty. Recently, in vitro studies have shown
that the nucleocytoplasmic traffic of human OREBP is a dynamic,
bi-directionally functional process. Whereas under isotonic conditions, OREBP
is detected in nuclear and cytoplasmic compartments, it accumulates
exclusively in the nucleus or cytoplasm when cells are subjected to hypertonic
or hypotonic challenges, respectively
(Tong et al., 2006). Previous
results from our group agree with these findings, since they suggest the
opposite pattern in the recruitment of putative OREBPs when rainbow trout
(Orozco et al., 2002b) and
tilapia (L.-B., unpublished observations) are transferred to seawater. These
results, together with new findings (Tong
et al., 2006), and the present results, support the proposal that
the functional motifs contained within the OREBP allow its bi-directional
shuttling in response to extracellular osmolarity. It should be noted that at
this stage, in contrast to mammals, our studies indicate that putative OREBP
translocates into the nucleus during hypo-osmolarity. Clearly, further studies
are necessary to fully elucidate the intimate mechanisms that regulate osmotic
homeostasis in the different species, especially in those continuously exposed
to environmental osmotic demands.
In contrast to the rapid osmolyte movement and the immediate gene
transcription that characterize the acute response to osmotic stress
(Pasantes-Morales et al.,
2006), in the present study the course of events extends for 12 h
after challenge. Thus, the time frame of this response is relatively slow, but
it coincides with the critical period for adaptation to a salinity change in
the killifish (Marshal et al.,
1999). Furthermore, within this time period, the present results
document a series of events that culminate in providing hepatocytes with
active TH. Indeed, both in vivo and in vitro experiments
showed a sequential correlation between the peak in translocation of putative
OREBP to the nucleus, the increase D2 transcription, and the subsequent rise
in enzymatic activity.
Recently, the mammalian liver has been recognized as an important
osmosensing and osmosignaling organ. Mammalian hepatocyte swelling or
shrinkage triggers an array of intracellular transduction signals that are
integrated with those that are hormone- and metabolic-dependant (for a review,
see Schliess and Haussinger,
2006). In elasmobranch osmoregulation, the liver is the main
provider of organic osmolytes including urea
(Hazon et al., 2003). The
osmoregulatory role of the teleostean liver has been less documented
(Fiess et al., 2007).
Osmoregulation is a highly expensive physiological process in terms of
metabolic energy. In fish it has been suggested to require from 20% to 50% of
the total energy expenditure, and is greater in freshwater than in seawater
(Boef and Payan, 2001;
Fiess et al., 2007). In
addition to supplying glucose and organic osmolytes
(Fiess et al., 2007), fish
liver contains the largest pool of glutamine, which is the major source of
ammonia as well as an important blood carrier for this nitrogenous waste
product (for reviews, see Wood,
1993; Haberle et al.,
2006). Two key enzymes involved in glutamine and ammonia
metabolism are glutamine synthetase and glutamate dehydrogenase, which
interestingly, at least in mammals, are both TH-dependent
(Doulabi et al., 2002). In this
context it is paradoxical that the current dogma considers the role of TH in
osmoregulation to be indirect. Indeed TH have been thought to support
long-term adaptive responses mediated by growth hormone, prolactin and
cortisol, among other classical osmoregulatory messengers
(Sakamoto and McCormick,
2006). However, previous studies from our laboratory
(Orozco et al., 1998;
Orozco et al., 2002b) and
recent studies in the seabream gill
(Klaren et al., 2007),
strongly suggest a more direct involvement of TH in the hydro-osmotic balance
in fish. Furthermore, our present results support the suggestion that a
putative hypo-osmotic, OREBP-mediated increase in hepatic D2 activity could be
an important endocrine component for the maintenance of hydro-osmotic
homeostasis in fish. Thus, we hypothesize that the local intra-hepatic
T3 increase that follows D2 activation during hypo-osmotic stress
may be instrumental for promote the hepatic synthesis of ammonia, providing
ammonium as a counter-ion for sodium absorption in the gill
(Salama et al., 1999).
In summary, this study examined the possible transcriptional regulatory role played by the ORE motifs in the 5' promoter region of FhDio2 during hypo-osmotic stress. Together, the present results provide strong evidence for the involvement of a putative OREBP kinase-dependant pathway as an important regulatory signal for FhDio2 transcription. To our knowledge, this is the first report that associates a response to hypo-osmotic stress with an ORE-regulated gene. The fact that this gene corresponds to the enzyme responsible for the local supply of intracellular T3 strongly suggests that, in addition to their permissive action, TH may play a more direct role in osmoregulatory homeostasis in fish, possibly by participating in hepatic ammonia metabolism. These findings may provide a clue to understanding the physiological function of TH in hydro-osmotic homeostasis in fish.
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Acknowledgments |
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