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First published online November 30, 2007
Journal of Experimental Biology 210, 4286-4297 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.009969
Endothelin and endothelin converting enzyme-1 in the fish gill: evolutionary and physiological perspectives
Department of Zoology, University of Florida, 221 Bartram Hall, Gainesville, FL 32608, USA and Mount Desert Island Biological Laboratory, Salisbury Cove, ME 04672, USA
* Author for correspondence (e-mail: khyndman{at}zoo.ufl.edu)
Accepted 24 September 2007
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Summary |
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Key words: endothelin, endothelin converting enzyme, Fundulus heteroclitus, killifish, gill
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Introduction |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionNote added in proofReferences |
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) and
natruresis in the kidney (Zeidel et al.,
1989). Endothelins are translated as 
200 amino acid (aa)
preproendothelins (preproEDN) that are initially cleaved by a furin-like
enzyme (Yanagisawa et al.,
1988) to form the relatively inactive 38 aa proendothelin (proEDN,
also known as Big-EDN) (Kimura et al.,
1989). Proendothelin is further cleaved to form the active 21 aa
EDN by the endothelin converting enzyme (ECE-1 and/or ECE-2)
(Shimada et al., 1994;
Xu et al., 1994). In mammals,
actions of EDNs are mediated via two G-protein coupled receptors:
endothelin A receptor (EDNRA), which preferentially binds EDN1
(Arai et al., 1990), and
endothelin B1 receptor (EDNRB1), which binds all three EDNs with equal
affinity (Sakurai et al.,
1990). In non-mammalian vertebrates, EDNs also equally bind to a
third G-protein coupled receptor, endothelin B2 receptor (EDNRB2)
(Lecoin et al., 1998), and in
amphibians EDN3 binds to an amphibian-specific receptor termed endothelin C
receptor (EDNRC) (Karne et al.,
1993).
Preproendothelin genes have been found in all major gnthastome clades
(Fig. 1), and there is evidence
for EDN1 regulation of vascular tone in fishes
(Olson et al., 1991;
Evans et al., 1996;
Evans, 2001;
Evans and Harrie, 2001;
Wang et al., 2001). In
addition, EDN1 inhibition of transport by the multidrug resistance-association
protein was demonstrated in shark (Squalus acanthias) rectal tubules
(Miller et al., 2002) and
killifish (Fundulus heteroclitus) renal tubules
(Masereeuw et al., 2000).
Recently, Evans et al. (Evans et al.,
2004) determined that exogenous (mammalian) EDN1 inhibited net
chloride transport in the killifish opercular epithelium, a tissue used as a
model for the SW teleost gill (Karnaky et
al., 1977). In teleosts, the gill is the main site for ion
balance, nitrogen excretion, acid–base regulation and gas exchange
(Evans et al., 2005).
Estuarine euryhaline fishes like the killifish (Fundulus
heteroclitus) encounter varying environmental salinities throughout the
day (Marshall, 2003),
resulting in a net gain or loss of ions depending on the water salinity; thus
the regulation of gill ion transport is an important mechanism to maintain
ionic homeostasis. Evans et al. hypothesized that EDN1 signaling cascades in
the gill may be a local regulator of ion balance in fishes
(Evans et al., 2004). Thus,
the purpose of this study was to determine if EDN1 and ECE1 are produced in
the killifish, and secondarily to determine if environmental salinity
regulates gill EDN1 and/or ECE mRNA expression. We were also
interested in determining the phylogenetic/evolutionary relationships of the
EDNs and ECE family of proteins.
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Materials and methods |
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EDN1 cDNA
All protocols and procedures were approved by the Institutional Animal Care
and Use Committee at the University of Florida. Previously published molecular
protocols were used (Hyndman et al.,
2006). Killifish were decapitated, and the gills of the right side
were removed and snap frozen in liquid nitrogen. Total RNA was then isolated
with TRI-reagent (Sigma, St Louis, MO, USA), and 5' and 3' RACE
cDNA was synthesized from 4 µg of total RNA using a GeneRacerTM Kit
(Invitrogen, Carlsbad, CA, USA) according to the manufacturer's protocols.
Published degenerate reverse EDN1 primers
(Wang et al., 2006) were used
in our initial 5' Touchdown RACE PCR following Invitrogen's protocols.
The polymerase used was 0.625 Units of Ex Taq, hot start, DNA polymerase
(Takara Bio, Madison, WI, USA) and the reactions were run in an Express
thermocycler (ThermoHybaid, Franklin, MA, USA). The PCR parameters were:
94°C for 2 min, 5 cycles of 94°C for 30 s, 72°C for 30 s, 5 cycles
of 94°C for 30 s, 70°C for 30 s, 30 cycles of 94°C for 30 s,
45°C for 30 s, 72°C 30 s, and a final 72°C for 5 min. PCR products
were visualized by ethidium bromide staining in 1.5% agarose gels, ligated
into pCR®4-TOPO vectors, and transformed into TOP10 chemically competent
cells using a TOPO TA Cloning® Kit for sequencing (Invitrogen). Plasmid
DNA was then sequenced in both directions at the Marine DNA Sequencing
Facility at the Mount Desert Island Biological Laboratory (Salisbury Cove, ME,
USA). Once we had the 5' end, specific killifish EDN1 primers
were designed (Table 1) and
3' Touchdown RACE PCR was performed to complete the cDNA sequences. The
PCR parameters were: 94°C for 2 min, 5 cycles of 94°C 30 s, 72°C
for 30 s, 5 cycles of 94°C for 30 s, 70°C for 1 min, 30 cycles of
94°C for 30 s, 50°C for 30 s, 72°C for 1 min, and a final 72°C
for 10 min. PCR products were cloned and sequenced as above.
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ECE cDNA
To obtain ECE cDNA, 4 µg of total gill RNA (extracted as
described above) was reverse transcribed using the First-strand cDNA
SuperscriptTM III reverse transcriptase kit (Invitrogen) with oligo-dT
primers. Initial degenerate primers used to generate ECE were designed using
CODEHOP (Rose et al., 2003)
and are recorded in Table 1.
PCRs were run on 0.5 µl of the oligo-dT cDNA, with 0.625 units of Ex Taq,
hot start (Takara) and standard cycling parameters. PCR products were cloned
and sequenced as described above.
Sequence and phylogenetic analysis
Sequence results for each transcript were assembled with GeneTools software
(BioTools Inc., Edmonton, Alberta, Canada) and killifish EDN1 and
ECE1 nucleotide sequences were searched for open reading frames
(ORFs). The resulting amino acid translations were analyzed with the basic
local alignment search tool (Blast) on the National Center for Biotechnology
Information (NCBI) website. The predicted amino acid sequences were aligned
with other full-length vertebrate EDN or ECE proteins using Clustal X
(Chenna et al., 2003). All
sequences were taken from GenBank or the Genome projects in Ensembl (e:!44
April 2007). Preproendothelin-1 sequences from each major vertebrate clade
(mammals to teleosts) were separately aligned, and similarities among the
sequences highlighted with GeneDoc (available at
http://www.psc.edu/biomed/genedoc),
including the expected cleavage sites for furin and ECE
(Yanagisawa et al., 1988;
Opgenorth et al., 1992). To
determine the relationship among our sequences and those from other organisms,
EDN and ECE alignments were exported to PHYML
(Guindon et al., 2005) and a
Fast Maximum-Likelihood test was performed, following the WAG model of amino
acid substitutions (Whelan and Goldman,
2001) and a calculated gamma of 1.023 and 1.03, respectively.
Branches were then tested for statistical significance by bootstrapping with
500 replicates.
Multiple tissue semi-quantitative PCR
To determine the distribution of EDN1A, EDN1B and ECE1
mRNA among tissues, relative duplexing semi-quantitative PCR was performed on
total RNA from gill, opercular membrane, brain, heart, stomach, intestine and
kidney tissue, as described previously
(Choe et al., 2004;
Choe et al., 2005). Briefly,
cDNA was produced from the tissues of a SW killifish as described above, but
random hexamer primers (not oligo-dT primers) were used so that ribosomal and
messenger RNA would be reverse transcribed. Non-degenerate primer pairs were
designed to amplify a product with high efficiency (e.g. high melting
temperature), and to minimize the chance of amplifying contaminating genomic
DNA, the primer pair was designed to include at least one intron–exon
boundary when possible (Table
1). A QuantumRNATM 18S internal standard primer kit (Ambion,
Woodward Austin, TX, USA) was used to control for variability in cDNA quality
and quantity between the different tissues tested. Duplexing PCR with primers
for 18S and either EDN1A, EDN1B or ECE1 were then optimized
to ensure that the reactions were terminated during the exponential phase.
Lastly, the products were visualized by ethidium bromide staining in 1.5%
agarose gels and digitized using the Biorad Gel DocTM XR System.
Salinity challenges
Killifish were acclimated to SW (approximate concentrations in mmol
l–1: Na+ 517, Ca2+ 9, K+ 12,
Cl– 486) (Choe and Evans,
2003) or freshwater (FW; Gainesville dechlorinated tapwater,
approximate concentrations in mmol l–1: Na+ 4,
Ca2+ 1, K+ 0.03, Cl– 0.40)
(Choe and Evans, 2003) for 2
weeks, at which point the SW killifish were transferred into FW (SW
FW)
and the FW killifish were transferred into SW (FWSW). An additional set
of killifish were removed and replaced into SW or FW as sham controls
(SWSW and FWFW, respectively). Immediately after transfer, 5 or 6
killifish from each treatment were sacrificed, the gills excised and snap
frozen for RNA extraction and cDNA synthesis. Killifish (N=5 or
6/treatment) were further sacrificed at 3, 8 and 24 h post transfer (acute
acclimations), as well as 30 days post transfer (chronic acclimation). RNA was
extracted from all of the samples and oligo-dT cDNA synthesized as described
above.
Quantitative real-time PCR
To determine the effects of environmental salinity on killifish gill
EDN1A, EDN1B and ECE1 mRNA levels, quantitative real-time
PCR (qRT-PCR) was performed. Nondegenerate primers were designed to amplify a
product between 50–100 bp across a predicted intron–exon boundary
(Table 1). L8 was used as an
internal control gene, as previously described
(Choe et al., 2005;
Choe et al., 2006). Each
sample was run in triplicate using 2 µl of 1/10 diluted original cDNA, 7.4
pmol of primers and SYBR® Green Master Mix (Applied Biosystems, Foster
City, CA, USA) in a total volume of 25 µl. The cycling parameters used
were: an initial denaturing step of 95°C for 10 min, 40 cycles of 95°C
for 35 s, 60°C for 30 s and 72°C for 30 s, followed by a melting curve
analysis to ensure only one product was amplified. Random samples were also
sequenced following qRT-PCR, confirming amplification of the target of
interest. To determine the degree of possible genomic contamination, qRT-PCR
was run using RNA samples that were not reverse transcribed, and we determined
that there was no genomic contamination. All qRT-PCRs were run on a MyiQ
quantitative thermocycler (Biorad, Hercules, CA, USA).
Each primer pairs' efficiency was determined by performing a tenfold
dilution curve using plasmid cDNA. Efficiency (E) for each primer pair was
calculated using the equation: E=–1+10(–1/slope), where
`slope' was the slope of the dilution curve. Each cycle threshold (CT) value
was subtracted from a randomly chosen control sample resulting in a 
CT,
and were analyzed using the Pfaffl equation
(Pfaffl, 2001): ratio=
ECTtarget/ECTL8. Each Pfaffl ratio was
then standardized to the average chronic seawater Pfaffl ratio.
Statistics
Values are expressed as means ± s.e.m. (standard error). For qRT-PCR
data, a two-factor ANOVA was performed to determine whether effect of
environmental salinity over time differed between SWFW transfers and
SWSW shams or FWSW transfers and FWFW shams. If statistical
significance was found a one-factor ANOVA was run to determine the effect of
time over a treatment group. Finally, all time points were compared to sham
time points with unpaired t-tests to determine if salinity transfers
altered mRNA expression. All values that did not meet homogeneity or equal
variance tests were log transformed to meet the assumptions of the ANOVA;
P=0.05.
Tissue preparation for in situ hybridization and immunohistochemistry
Killifish gills were fixed in 4% paraformaldehyde in 10 mmol
l–1 phosphate buffered saline (PBS) pH 7.3, for 24 h,
dehydrated in an increasing concentration of ethanol, cleared in Citrisolv
(Fisher Scientific, Pittsburgh, PA, USA), and embedded in paraffin wax. The
tissue blocks were cut at 7 µm, placed on Superfrost Plus slides (Fisher
Scientific), and heated at 37°C overnight.
In situ hybridization
mRNA for EDN1A, EDN1B and NKA
(Na+,K+-ATPase) mRNA were visualized using in
situ hybridization. An ECE1 mRNA probe was not made because our
partial sequence of that transcript was from the middle of the sequence, a
region that in other fishes is >65% identical to ECE2, and we were
afraid of the potential cross-reactivity of this probe. Specific digoxigenin
(DIG)-RNA probes (sense and antisense) were made against the 3' end of
the transcripts [including untranslated regions (UTRs) for the EDNs;
these regions were <60% identical]. For EDN1A the probe was made
from position 520 to the end of the transcript, including the polyA tail (420
bp long). The EDN1B probe was made from position 515 up to and
including the polyA tail (419 bp). Both of these transcripts were cloned as
described above. A killifish NKA mRNA probe was also made based upon
the complete killifish NKA sequence (AY057072). The probe was made from base
pairs 915–3115. This transcript was also cloned and sequenced to ensure
it was indeed NKA. All of the transcripts were linearized by T3/T7
PCR amplification from the plasmids containing the sequences of interest.
DIG-RNA probes were generated by incubating 100–200 ng of the linearized
transcripts with the DIG RNA Labeling mix (Roche Applied Science,
Indianapolis, IN, USA) following the manufacturer's protocols, at 37°C for
16 h followed by treatment with DNAse for 1 h at 37°C. The DIG-RNA probes
were purified using mini Quick Spin RNA columns (Roche) following the
manufacturer's instructions, eluted in 80 µl of diethyl pyrocarbonate
(DEPC) treated water and stored at –80°C until use.
To determine which gill cells expressed EDN1A or EDN1B mRNA, gill tissue slides were rehydrated in two changes of Citrisolv, followed by incubation in a series of decreasing concentration of ethanol washes. The slides were placed in sterile 10 mmol l–1 PBS and post-fixed in 4% PFA for 10 min at room temperature (25°C). Following this, the slides were rinsed in sterile 10 mmol l–1 PBS and incubated in proteinase K (5 mg ml–1) at room temperature for 5 min. Again, they were washed in 10 mmol l–1 PBS and post fixed in 4% PFA for 10 min to inactivate the proteinase K. After a final PBS wash, the slides were incubated in prehybridization solution (50% formamide, 10% dextran sulphate, 2% blocking reagent, 0.1% CHAPS, 1% Tween 20, 5 mmol l–1 EDTA, pH 8.0, 5x SSC, 50 µg ml–1 heparin, 1 mg ml–1 tRNA, in DEPC-water) for 2 h at room temperature. Next 200–500 ng of DIG-RNA probes were added to fresh prehybridization solution and the slides were left to incubate at 60°C for 18–24 h. Following this, the tissues were washed for 30 min in 2x SSC at room temperature, 2x SSC at 60°C, two 0.2x SSC 60°C washes, one 0.2x SSC at room temperature and KTB (50 mmol l–1 Tris pH 7.5, 100 mmol l–1 NaCl and 10 mmol l–1 KCl) at room temperature. The tissues were then blocked in 20% normal goat serum diluted in KTB for 1 h at room temperature and incubated in 7.5 U ml–1 of sheep anti-DIG-AP, Fab fragments (Roche) diluted in normal goat serum (NGS), overnight at 4°C. The slides were then washed in three changes of KTB and incubated in alkaline phosphatase buffer (100 mmol l–1 Tris, pH 9.5, 100 mmol l–1 NaCl, 50 mmol l–1 MgCl2) for 30 min at room temperature. Visualization of the probes was achieved by incubating the tissues in BCIP/NBT Substrate Kit, 5-bromo-4-chloro-3-indolyl phosphate/nitroblue tetrazolium (Vector Labs, Burlingame, CA, USA) with levamisole, following manufacturer's instructions, at room temperature until the signal developed (2–6 h). Images were captured using an Olympus BX60 light microscope with a Hitachi KP-D50 digital camera. Image contrast and brightness were adjusted with Photoshop CS (Adobe, San Jose, CA, USA).
Immunohistochemistry
Slides were analyzed following published methods
(Piermarini et al., 2002;
Hyndman et al., 2006). Slides
with chronic SW and FW acclimated killifish gill tissue were incubated in
primary antibodies: polyclonal, anti-human-proEDN1 (1/1000 dilution) (Phoenix
Pharmaceutical, Burlingame, CA, USA) made against the complete 38 aa of human
proEDN1, which is 74% identical to both killifish EDN1s. Monoclonal, anti-NKA
(
5, 1/1000) was developed by Dr Douglas Fambrough, and was obtained
from the Developmental Studies Hybridoma Bank, which was developed under the
auspices of the National Institute of Child Health and Human Development of
the University of Iowa, Department of Biological Sciences, Iowa City, IA
52242, USA.
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionNote added in proofReferences |
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We have sequenced 1696 bp from the middle of the killifish ECE1 cDNA (accession no. EU009476). This translates into 565 aa from the killifish ECE1. Endothelin converting enzymes are found in all organism including Bacteria and Archaea (Fig. 3). As seen in Fig. 3, the non-vertebrate ECEs are not well resolved (i.e. lancelet ECE groups with locust and sea urchin, while sea squirt groups with hydra), and although there is no clear explanation for this, Fig. 3 shows three distinct ECE clades: ECE1, ECE2 and non-vertebrate ECE. Our partial sequence of ECE1 from the killifish groups with the other fish ECE1 sequences confirming it is ECE1.
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Tissue distributions
Using duplexing relative semi-quantitative PCR we found EDN1A mRNA
in the gill, opercular epithelium, brain, heart, stomach, intestine and kidney
of the killifish (Fig. 4).
Relatively high expression was found in the gill, brain and kidney.
EDN1B mRNA was not found in the opercular epithelium, and had very
little expression in the gill, but was highly expressed in the brain, kidney
and intestine. Finally, ECE1 mRNA was found in all of the tissues
tested, with highest expression in the stomach, intestine and gill
(Fig. 4).
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; Marshall,
2003; Choe et al.,
2006; Hyndman et al.,
2006). In contrast, EDN1B mRNA expression was rare (as
was seen in the EDN1B tissue distribution described above); however,
it was found in some lamellar pillar cells
(Fig. 5D,F) and in epithelial
cells adjacent to the environment (Fig.
5D). Incubation of slides with sense probes produced a little
non-specific staining (Fig.
5B,E).
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;
Goniakowska-Witalinska et al.,
1995; Zaccone et al.,
1996; Mauceri et al.,
1999). This immunolocalization matches the killifish EDN1
mRNA localization shown in Fig.
5. Slides incubated in NGS and double labeled with NKA were done
as negative controls, and showed no non-specific staining of the proEDN1
secondary antibody and chromagen (Fig.
6B,D).
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FW) transfers
(Fig. 7A,C). ECE1 mRNA
increased four- and sixfold compared with sham ECE1 mRNA levels at 8
and 24 h post a FWSW transfer (Fig.
7E). EDN1A mRNA levels did not change significantly with
acute acclimation from SWFW compared to sham (SWSW) treatments
(Fig. 7B), but 24 h
EDN1B mRNA levels were almost threefold higher compared to sham
EDN1B mRNA levels at 24 h (Fig.
7D). ECE1 mRNA levels were twofold higher after 3 and 24
h of acclimation to FW but did not differ from sham values at 8 h post
transfer (Fig. 7F). With
chronic acclimation (30 days), there were no statistical differences between
the SW- and FW-acclimated killifish for EDN1A, EDN1B or ECE1
mRNA levels (Fig. 8). In
addition, preproEDN1 (protein) immunolocalization did not differ between fish
chronically acclimated to FW (Fig.
6A,B) or SW (Fig.
6C,D).
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Discussion |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionNote added in proofReferences |
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Gill expression of EDN1 mRNA and proEDN1 protein
In the killifish gill we found EDN1A mRNA expression in epithelial
cells of the interlamellar region, and EDN1B mRNA expression was
found in pillar cells, and in cells adjacent to the environment, in the
interlamellar region. Not only are these two transcripts expressed in
different levels within a tissue (Fig.
4), they are also expressed in different cells within the gill.
From our immunohistochemical experiments, we found proEDN1 immunoreactivity in
epithelial cells adjacent to the NKA immunoreactive cells. NKA is commonly
used as a marker for the ion transporting, MRCs of the fish gill
(Katoh et al., 2001;
Marshall, 2003;
Evans et al., 2005).
Proendothelin immunoreactivity was also found on pillar cells, which is in
agreement with our in situ hybridization findings. ProEDN was
immunolocalized to gill neuroendocrine cells (NECs) of the eel (Conger
congo), catfish (Heteropneustes fossilis) and dogfish
(Scyliorhfnus canicula), and the morphology of these cells matches
that of the proEDN immunoreactive epithelial cells in the killifish gill
(Zaccone et al., 1996).
Endothelin production in the pillar cells was suspected by earlier workers
(Sundin and Nilsson, 1998;
Stenslokken et al., 1999), who
showed that infusion of mammalian EDN1 into the lamellae of the rainbow trout
resulted in a `constriction of the vascular sheet' (of the lamellae)
and that this was likely due to constriction of the pillar cells. The authors
hypothesized that hormonal control of pillar cell tone may be one mechanism to
match respiratory needs of a fish while minimizing ion fluxes. There is no
evidence that pillar cells are innervated
(Bettex-Galland and Hughes,
1972; Bettex-Galland and
Hughes, 1973); thus endocrine/paracrine/autocrine signaling
molecules may be the regulators of pillar cell tone. Pillar cells contain
contracting filamentous material
(Bettex-Galland and Hughes,
1972; Bettex-Galland and
Hughes, 1973) and recently an actin-binding protein was described,
FHL5, that is highly expressed in these cells, suggesting that they are
capable of contraction (Mistry et al.,
2004). Video microscopy was used to demonstrate in vivo
that pillar cells do contract with EDN1 infusion
(Stenslokken et al., 1999).
Recently, EDN receptors EDNRA and EDNRB were immunolocalized in the fish gill,
and EDNRB was found throughout the gill vasculature, NECs and pillar cells of
the cod Gadus morhua (Stenslokken
et al., 2006). EDNRA was described in nerve fibers running along
the length of the filament and innervating the gill vasculature
(Stenslokken et al., 2006). In
the fugu Takifugu rubripes EDNRA was found on the pillar cells and in
erythrocytes (Sultana et al.,
2007). Studies from our lab in the killifish have found EDNRB
receptors throughout the gill vasculature and pillar cells, and EDNRA
receptors on the mitochondrion-rich cell (K.A.H. and D.H.E., unpublished
observations). In the long horn sculpin Myoxocephalus
octodecimspinosus, EDNRA receptors were found on the pillar cells, while
EDNRB was found throughout the gill vasculature (K.A.H. and D.H.E.,
unpublished observations). Evidently, there is species-specific EDN receptor
distribution in the gill of fishes.
Acute and chronic salinity acclimations
Killifish usually live in estuaries where there are rapid changes in
environmental conditions such as salinity and temperature
(Marshall, 2003). We tested
the effects of rapid changes of environmental salinity on mRNA expression of
EDN1A, EDN1B and ECE1. EDN1 transcript levels did not change
with chronic acclimation to FW or SW (Fig.
8). In addition, proEDN1 immunoreactivity in the gill did not
differ between SW and FW acclimated killifish
(Fig. 6). However,
EDN1B and ECE1 mRNA levels increase with acute FW
acclimation, suggesting that more active EDN1 protein is produced.
10–8 mol l–1 mammalian EDN1 can inhibit net
chloride transport in the killifish opercular epithelium, and this is
predominately due to stimulation of cyclo-oxygenase (COX) and subsequent
prostaglandin production (Evans et al.,
2004). These findings suggest that during transfer to a
hypo-osmotic environment, EDN1B and ECE1 protein levels increase, resulting in
an increase in active EDN1 that could potentially inhibit net chloride
transport, helping the fish retain ions. However, we cannot rule out that EDN1
signaling in the gill is different than what was described in the killifish
operculum (Evans et al., 2004)
(see below). In addition, it is undetermined how volume stress, such as occurs
during a rapid change to a hypo-osmotic environment, effects blood flow
through the gill. EDN1B was found on gill pillar cells, and may play
a role in regulating blood flow during blood volume increases; however, this
is an unexplored area of fish gill physiology.
Although we found an increase in EDN1 during acclimation to FW, we
unexpectedly found a sixfold increase in ECE1 mRNA levels with acute
SW acclimation, suggesting that there is an increase in ECE1 production during
this period. This in turn would result in more EDN1 production because the
proteolytic cleavage of proEDN1 to EDN1 by ECE1 is a rate-limiting step
(D'Orleans-Juste et al.,
2003). Our attempts to measure EDN1 production in the fish gill by
enzyme immunoassay and Tris-Tricine western blotting were unsuccessful, but
measurements of EDN1 levels are necessary to fully understand the role of EDN1
cell signaling in the fish gill. Recently, a 3.4-fold increase was shown in
COX-2 mRNA levels in the killifish gill
(Choe et al., 2006), 3 h post
a FWSW and a 2.6-fold increase 3 h post a SWFW transfer, and the
authors hypothesized that the increase in COX-2 is an important mechanism for
gill cell survival during osmotic stress. A similar result has been
demonstrated in mammalian medullary interstitial cells (that experience large
changes in osmotic stress), which require functional COX-2 to survive
(Hao et al., 1999;
Hao et al., 2000). Medullary
interstitial cells also contain EDNRs but do not produce EDN1
(Dean et al., 1996).
Endothelin has been shown to stimulate COX-2 in a variety of mammalian tissues
(Hughes et al., 1995;
Chen et al., 2003), and EDN1
signaling via endothelial ENDRB results in the production of
prostacyclin (Warner et al.,
1989; Hirata et al.,
1993). Taken together, our findings suggest that during rapid
changes in environmental salinity, gill cell survival during this osmotic
stress may be accomplished by increased EDN1 production and subsequent
stimulation of COX production of prostaglandins. To the best of our knowledge,
it is unclear what aids cell survival during salt or water load in fishes, and
it is plausible, since EDN1 and ECE1 are ubiquitously expressed, that this may
be a more global change in their signaling patterns and is not a gill-specific
phenomenon; however, this is yet to be determined and experiments testing
these hypotheses are needed. In addition, studies blocking aspects of EDN1
signaling in the gill and subjecting these fish to salinity challenges are
vital in understanding EDN1 function during osmotic stress. This technique has
been successful in mice models, where kidney collecting duct EDN1 (or EDNRB1)
knockout mice who are fed a high salt diet are unable to excrete the excess
Na+ accumulated and are severely hypertensive
(Ahn et al., 2004;
Ge et al., 2006), suggesting
that EDN1 is necessary for salt excretion in mammals. Applications of these
types of techniques to fish models are necessary to fully understand the
in vivo role of EDN signaling in the fish gill.
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). Physiological responses to EDN1 have also been
demonstrated in the spiny dogfish shark
(Evans et al., 1996;
Evans and Gunderson, 1999),
again suggesting that EDN1 is produced endogenously. Currently, the exact
evolutionary history of this family of peptides is not clear. EDN sequences
from hagfish, lamprey and sharks are necessary to determine when these
peptides arose, and to determine if it was due to gene/genome duplications or
other evolutionary events over vertebrate evolution. Interestingly, mutations
in EDN1, endothelin receptors (EDNRs) or ECE result in severe craniofacial
developmental abnormalities, and these phenotypes are often lethal
(Kurihara et al., 1994;
Brand et al., 1998;
Clouthier and Schilling, 2004;
Nair et al., 2007), suggesting
the EDN signaling is necessary for development in vertebrates, and it may be a
key innovation in the radiation of vertebrates.
Unlike EDNs, which are only found in vertebrates, ECE is found in all
organisms, including Bacteria and Archaea. In
Fig. 3, our maximum likelihood
analyses reveal three distinct groups of ECEs: prokaryote, fungal and
invertebrate ECE, vertebrate ECE1 and vertebrate ECE2. This suggests a gene
duplication event sometime after the chordate–vertebrate split, but
before the teleost radiation. Since there is no molecular evidence for EDNs or
EDNRs in animals basal to the vertebrates why would they have an ECE?
Endothelin converting enzymes are zinc-dependant metalloendoproteases and part
of the Neprelysin and Kell family (Shimada
et al., 1994; Xu et al.,
1994). In vertebrates, ECE can function as a monomer or dimer;
however, for effective proteolytic cleavage of proEDN1 to EDN1, dimerization
at Cys412 is preferential
(Shimada et al., 1996). In
contrast, hydra ECE (Zhang et al.,
2001) and the other invertebrate, fungal and prokaryote ECEs are
missing Cys412 and are believed to function as monomers
(Zhang et al., 2001).
Vertebrate ECE has been shown to cleave peptides other than proEDN, including
bradykinin, angiotensin I and substance P
(Hoang and Turner, 1997;
Johnson et al., 1999),
suggesting that ECE may be a generalist protease. Although the native
substrates cleaved by ECE in non-vertebrate organisms are undetermined, it is
plausible that ECE originally cleaved substrates found in all organisms, and
during vertebrate evolution started functioning as a dimer and preferentially
cleaving proEDN.
Tentative model for EDN1 signaling in the killifish gill
To summarize our findings, we propose the following model
(Fig. 9) of paracrine and
autocrine EDN1 signaling in the fish gill. The diagram shows a lamellar
cross-section of the gill (same orientation as the gills in Figs
5,
6,
7), with pillar cells (PCs)
highlighted in grey and adjacent pavement cells (PVCs) in white. In the
intralemallar region there are two MRCs and an NEC above the gill vasculature.
Cyclo-oxygenase-2 (COX-2) and neuronal nitric oxide synthase (nNOS) were
previously immunolocalized in the killifish gill, to MRCs
(Choe et al., 2006) and NECs,
nerve fibers and lamellar arterioles (LA)
(Hyndman et al., 2006),
respectively. NKA was immunolocalized to the basolateral membrane of the MRC
(see Katoh et al., 2001;
Choe et al., 2006;
Hyndman et al., 2006) and the
chloride channel, cystic fibrosis transmembrane conductance regulator (CFTR),
to the apical membrane of the MRC (Katoh
et al., 2001). From our studies and others, EDNRB were found
throughout the gill vasculature (K.A.H. and D.H.E., unpublished)
(Stenslokken et al., 2006),
and EDNRB and EDNRA were on the pillar cells, depending on the species (K.A.H.
and D.H.E., unpublished observations)
(Stenslokken et al., 2006;
Sultana et al., 2007). EDNRA
were found on MRCs in the killifish gill (K.A.H. and D.H.E., unpublished
observations). Here we present EDN1 expression in cells adjacent to the MRC
(likely NECs) and pillar cells. This suggests a paracrine role of EDN1
signaling, given that it is produced in the NEC and can bind to receptors on
the adjacent MRCs (Fig. 9,
pathway 1) where it potentially stimulates COX-2 activity, resulting in cell
survival during osmotic stress and/or alter ion transport by the MRC as
previously hypothesized (Evans et al.,
2004). EDN1 can also potentially act as a paracrine binding to
EDNRB receptors on the gill vasculature and lamellar arterioles, suggesting it
can regulate perfusion of the lamellae
(Fig. 9, pathway 1). It also
can act as an autocrine on the pillar cells, further supporting the role of
regulation of local perfusion across a lamella to meet the respiratory needs
of the fish (Fig. 9, pathway 2)
(Sundin and Nilsson, 1998;
Stenslokken et al., 1999). It
may also help maintain lamella integrity during rapid increases in plasma
volume during exposure to a hypo-osmotic environment. This is the first model
to depict EDN1 signaling in the fish gill, and in the future, studies
determining the specific function of EDN1 in the gill and whole fish, are
necessary to understand its role in normal fish physiology.
List of abbreviations and symbols
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) are indicated. Asterisks mark every
20th amino acid. (B) Predicted active EDN1 structure from the killifish,
modeled after Webb (Webb,
1997
