Short-term transformation and long-term replacement of branchial chloride cells in killifish transferred from seawater to freshwater, revealed by morphofunctional observations and a newly established `time-differential double fluorescent staining' technique
Department of Aquatic Bioscience, Graduate School of Agricultural and Life Sciences, University of Tokyo, Tokyo 113-8657, Japan
* Author for correspondence (e-mail: fkatoh{at}marine.fs.a.u-tokyo.ac.jp)
Accepted 11 August 2003
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Summary |
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Key words: branchial chloride cell, killifish, Fundulus heteroclitus, time-differential double fluorescent staining
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Introduction |
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Recent studies have shown that chloride cells in some euryhaline fishes
alternate their morphology and ion-transporting functions to meet abrupt
environmental osmotic changes. In the tilapia yolk-sac membrane, a sequential
observation technique revealed that single freshwater-type chloride cells were
transformed into seawater-type multicellular complexes during seawater
adaptation (Hiroi et al.,
1999). Sakamoto et al.
(2000
) also showed that
mudskipper Periophthalmus modestus closed the apical pit of chloride
cells following transfer from seawater to freshwater. In seawater-adapted
killifish, the cystic fibrosis transmembrane conductance regulator (CFTR) and
Na+,K+,2Cl- cotransporter have been
demonstrated in the apical pit and cytoplasm of MR cells, respectively. These
ion transporters are redistributed during freshwater adaptation
(Marshall et al., 2002
).
In most teleosts examined so far, chloride cells became larger and denser
when the fish were transferred from freshwater to seawater, concomitant with
increases in gill Na+/K+-ATPase activity
(Langdon and Thorpe, 1985;
Richman et al., 1987
;
McCormick, 1995
; Uchida et
al., 1996
,
2000
;
Shiraishi et al., 1997
). This
ion-transporting enzyme is located in the tubular system that is continuous
with the basolateral membrane, and plays a key role in ion-transporting
functions of chloride cells (McCormick,
1995
). Thus, chloride cell morphology and gill
Na+/K+-ATPase activity were thought to be reliable
indices of seawater adaptability. However, this is not the case with killifish
Fundulus heteroclitus, which also exhibits morphologically distinct
freshwater- and seawater-type chloride cells in the respective media
(Marshall et al., 1997
;
Katoh et al., 2001
). In
killifish, chloride cells are mostly located in a flat region of the
afferent-vascular edge (the trailing edge, in terms of water flow) of the gill
filament, and the chloride cell size is generally larger in freshwater-adapted
than in seawater-adapted fish. Furthermore, gill
Na+/K+-ATPase activity, as well as oxygen consumption,
does not differ between freshwater- and seawater-adapted fish
(Marshall et al., 1999
;
Katoh et al., 2001
),
suggesting that both types of chloride cells are equally active in the two
environments but exhibit different ion-transporting functions.
According to the current model for transcellular ion transport by chloride
cells, various ion-transporting proteins are placed in either the apical or
basolateral membrane. In seawater-type chloride cells, for example,
Cl- enters the cell, together with Na+ and
K+, across the basolateral membrane via the
Na+,K+,2Cl- cotransporter, and accumulates
intracellularly so that Cl- exit occurs down its electrochemical
gradient through Cl- channels in the apical membrane. The driving
force for Cl- secretion is the Na+ electrochemical
gradient established by Na+/K+-ATPase in the basolateral
membrane. Meanwhile, Na+ secretion occurs down its electrochemical
gradient via a cation-selective paracellular pathway. In killifish,
the CFTR has been identified as the apically located Cl- channel
(Marshall et al., 1995; Singer
et al., 1988), and thus the existence of CFTR in the apical membrane could
provide evidence for functional seawater-type chloride cells.
In seawater-type chloride cells in the killifish gills, the apical membrane invaginates to form a pit, and the cells often interdigitate with neighboring accessory cells. In freshwater-type cells, on the other hand, the apical membrane is flat or protrusive, being equipped with microvilli. Although the structural difference is evident between freshwater- and seawater-type chloride cells in killifish, it is not clear whether one cell type is replaced by another cell type, or whether one cell type changes its function and morphology into another type, following transfer between freshwater and seawater.
In this study, we aimed to clarify both the short- and long-term responses of gill chloride cells to direct transfer from seawater to freshwater in killifish. To visualize the chloride cell replacement, or cell turnover, following freshwater transfer, we developed a `time-differential double fluorescent staining (TDS) technique', in which in vivo labeling of chloride cells was performed on two separate days, using two distinguishable mitochondria-specific fluorescent probes. Our findings revealed functional and morphological transformation of chloride cells from seawater- to freshwater-type in a short term, which was followed by recruitment of newly differentiated, freshwater-type cells as a long-term response to freshwater transfer.
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Materials and methods |
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Time-course observations following transfer from seawater to
freshwater
Forty seawater-acclimated fish weighing 2.8-7.8 g were transferred directly
to freshwater in 40 liter tanks at 25°C, and removed at 0 h (initial
controls), 3 h, 12 h, 1 day, 3 days, 7 days, 14 days and 30 days after
transfer. The fish (N=5 each) were anesthetized with 0.05%
2-phenoxyethanol, and blood was collected from the caudal vessels into
capillary tubes. The plasma was separated by centrifugation at 4000
g for 5 min. Plasma Na+ concentrations were
measured using an atomic absorption spectrophotometer (Hitachi Z-5300, Japan).
The gills were removed and fixed in 4% paraformaldehyde (PFA) in 0.1 mol
l-1 phosphate buffer (PB, pH 7.4) for 24 h for whole-mount
immunocytochemistry, and in 2% PFA, 0.2% glutaraldehyde (GA) in the same
buffer for 3 h for CFTR immunocytochemistry. For transmission (TEM) and
scanning (SEM) electron microscopy, the gills were fixed in 2% PFA, 2% GA in
PB for 24 h. The fixed gill samples were stored in 70% ethanol. For
histological observations, we examined three animals in each experimental
group.
Antibodies
For the detection of chloride cells in the gill filaments, we used an
antibody specific for Na+/K+-ATPase. The antiserum
(NAK121) was raised in a rabbit against a synthetic peptide corresponding to
part of the highly conserved region of the Na+/K+-ATPase
-subunit, and the specific antibody was affinity-purified
(Katoh et al., 2000
;
Uchida et al., 2000
). The
specificity was confirmed by western blot analysis
(Katoh et al., 2000
).
The antibody used to detect killifish CFTR was a mouse monoclonal antibody
to human CFTR (R&D Systems, MN, USA). This antibody was raised against a
carboxy-terminal sequence of human CFTR, which was identical to that of
killifish CFTR (Singer et al.,
1998).
Western blot analysis for antibody to CFTR
Since the CFTR content in the gill was thought to be too small for
detection by western-blot analysis, the sample was subjected to
immunoprecipitation prior to western blotting. The gill filaments were scraped
in 1 ml of lysis buffer consisting of IP buffer (pH 7.4; 140 mmol
l-1 NaCl, 2 mmol l-1 KCl, 10 mmol l-1 Hepes
and 5 mmol l-1 EDTA), inhibitors (10 mmol l-1
benzamidine, 1 µg ml-1 Pepstatin A and 2 mmol l-1
phenyl methyl sulfonyl fluoride) and 1% Triton-X 100, and left on ice for 20
min to lyse the cells. The lysate was centrifuged at 5000 g
for 5 min at 4°C, and the supernatant was incubated with 1 µl of the
CFTR antibody at room temperature for 3 h. Slurry (20 µl) containing 50%
protein A sepharose beads (Amersham Biosciences, Uppsala, Sweden) blocked
overnight with 1% bovine serum albumin (BSA), was added to the sample, and the
mixture was incubated for 1 h at 4°C. After washing five times with IP
buffer containing inhibitors and centrifugation at 10 000 g
for 30 s, 30 µl of hot Laemmli buffer
(Laemmli, 1970) containing 5%
ß-mercaptoethanol was added to beads binding to the antibody, and the
mixture was incubated at 65°C for 15 min. The sample was centrifuged at 10
000 g for 2 min, and the supernatant was frozen for later
western blotting. The supernatant was separated by SDS-polyacrylamide gel
electrophoresis using 7.5% polyacrylamide gels (Atto, Japan). After
electrophoresis, the protein was transferred from the gel to a polyvinylidene
difluoride membrane (Atto). The specific protein band on the membrane was
detected with anti-CFTR diluted 1:100, according to the method described in
Katoh et al. (2000
).
Whole-mount immunocytochemistry
The gill filaments were removed from gill samples fixed in 4% PFA. After
washing in 0.01 mol l-1 phosphate-buffered saline (PBS, pH 7.4)
containing 0.05% Triton X-100, whole-mount preparations of the gill filaments
were incubated overnight at 4°C with a mixture of anti-human CFTR at a
final dilution of 1:1000 and anti-Na+/K+-ATPase (NAK121)
labeled with Alexa Fluor 546 (Molecular Probes, OR, USA) diluted 1:500 with
PBS containing 0.05% Triton X-100, 10% normal goat serum (NGS), 0.1% BSA,
0.02% keyhole limpet hemocyanin (KLH) and 0.01% sodium azide. The samples were
washed for 1 h with PBS containing 0.05% Triton X-100, and then incubated with
anti-mouse rabbit IgG labeled with Alexa Fluor 488 (Molecular Probes)
overnight at 4°C. The samples were then washed for 1 h with PBS, placed in
a chamber slide closed with a coverslip, and observed with a confocal laser
scanning microscope (LSM 510, Zeiss, Oberkochen, Germany). The 488 nm
argon-ion laser was used for Alexa Fluor 488 and the 543 nm helium-neon laser
for Alexa Fluor 546, to give the appropriate excitation wavelengths. The sizes
of chloride cells stained with Alexa Fluor 546-labeled
anti-Na+/K+-ATPase were measured on stored LSM images by
means of an internal program. The chloride cell area was obtained from 20
cells per gill filament. Three gill filaments were examined per individual
(N=3).
Fluorescence microscopy
The gills fixed in 2% PFA, 0.2% GA in PB were immersed in 30% sucrose in
PBS for 1 h, and embedded in Tissue-Tek OCT compound (Sakura Finetek, Japan)
at -20°C. Cryosections (2 µm thick) were cut on a cryostat (CM 1100,
Leica, Germany) at -20°C, and collected onto amino propyltriethoxy silane
(APS)-coated slides (Matsunami, Japan). The sections were then incubated with
a mixture of anti-human CFTR at a final dilution of 1:1000 and
anti-Na+/K+-ATPase (NAK121) labeled with Alexa Fluor 546
diluted 1:1000 with PBS containing 2% NGS, 0.1% BSA, 0.02% KLH and 0.01%
sodium azide (NB-PBS) overnight at 4°C, and then with anti-mouse rabbit
IgG labeled with Alexa Fluor 488 for 2 h at room temperature. The sections
were observed under a fluorescence microscope with blue (excitation, 450-490
nm; emission, 520-560 nm) and green (excitation, 510-560 nm; emission, >590
nm) excitation filter blocks for Alexa Fluor 488 and for Alexa Fluor 546,
respectively (Nikon E800, Japan).
Scanning electron microscopy
Gill filaments fixed for electron microscopy were dehydrated in ethanol,
immersed in 2-methyl-2-propanol, and dried using a freeze-drying device (JEOL
JFD-300, Japan). Dried samples were mounted on specimen stubs, and coated with
platinum palladium in an ion sputter (Hitachi E-1030, Japan), before
examination using a SEM (Hitachi S-4500).
Transmission electron microscopy
Tissues fixed for electron microscopy were postfixed in 1% osmium tetroxide
in PB for 1 h at room temperature. After dehydration in ethanol, the gill
tissues were transferred to propylene oxide and embedded in Spurr's resin.
Ultrathin sections were cut with a diamond knife, mounted on grids, stained
with uranyl acetate and lead citrate, and observed using a TEM (Hitachi
H-7100).
Chloride cell replacement during freshwater adaptation
To examine chloride cell replacement in response to salinity change,
seawater-acclimated killifish weighing 15.8-27.8 gwere transferred to
freshwater. For the purpose of comparison, the fish of similar size (19.0-36.9
g) acclimated to seawater or freshwater were also examined for chloride cell
replacement under constant environmental salinity. Experimental fish were
fin-clipped for individual discrimination. Chloride cell replacement was
examined using a newly developed time-differential double fluorescent staining
technique (see below). During the experiment, the fish were kept in the dark
to prevent the fluorescent probes from fading. The water temperature was
maintained at 25°C throughout the experiment.
Time-differential double fluorescent staining
To examine chloride cell replacement, we developed a time-differential
double fluorescent staining (TDS) technique, in which in vivo vital
staining for chloride cells was performed just before transfer (day 0) and 3
days after transfer (day 3), using two distinguishable mitochondria-specific
fluorescent probes.
On day 0, the fish were immersed one by one in 200 ml of 50 µmol l-1 Rhodamine 123 (Molecular Probes) dissolved in the respective environmental waters for 3 h to label chloride cells (pre-existing chloride cells). To assess the chloride density on day 0, biopsy samples of gill filaments were removed immediately after the first labeling under anesthesia with 0.05% 2-phenoxyethanol. The samples were placed in a chamber slide with PBS topped by a coverslip, and observed under a fluorescence microscope with the blue-excitation filter block (excitation, 450-490 nm; emission, 520-560 nm; Nikon). The fish were allowed to recover in their respective environmental waters for 30 min after the biopsy sampling, and then transferred from seawater to freshwater, whereas those in control groups were maintained in their respective environments throughout the experiment. On day 3, for the second labeling, the fish were incubated for 3 h as described above, in 1 µmol l-1 MitoTracker Red CM-H2XRos (Molecular Probes) in their respective environmental waters. MitoTracker was first dissolved in dimethyl sulfoxide at a concentration of 1 mmol l-1, and then diluted to the final concentration with environmental waters. After 3 h incubation, gill filaments were removed again from the same individuals under anesthesia. The filament samples were prepared as stated above, and observed under the fluorescence microscope with blue- and green- (excitation, 540-580 nm; emission, 600-660 nm, Nikon) excitation filter blocks. Consequently, the pre-existing cells labeled on day 0 were recognized as Rhodamine 123-positive cells stained in green, and newly differentiated cells during the last 3 days, as well as the preexisting cells, were stained with MitoTracker in red on day 3. Therefore, newly differentiated cells were identified as Rhodamine 123-negative, MitoTracker-positive cells on day 3.
The fluorescence-microscopic images were recorded with a digital camera (DXM1200, Nikon) attached to the microscope. To determine the chloride cell turnover, the numbers of Rhodamine 123-positive cells on days 0 and 3, and MitoTracker-positive cells on day 3, were counted in the afferent-vascular edge of 15 filaments from 3 individuals in each experimental group.
Statistics
Data are presented as mean ± standard error of the mean
(S.E.M.). The significant differences in the plasma Na+
concentration and chloride cell size were determined by Games Howell's test
after analysis of variance (ANOVA) by Bartlett's test. Significant differences
in the density of Rhodamine 123-positive chloride cells (pre-existing cells)
between days 0 and 3 and the total cell density between days 0 and 3 were
examined using a two-sample t-test. The significance of difference in
frequencies of MitoTracker-positive cells (newly differentiated cells) on day
3 between three experimental groups was determined by the
2-test for independence after ANOVA among individuals in each
group.
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Results |
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Western blot analysis for CFTR
In western blot analysis, the anti-CFTR recognized one specific protein
band with a molecular mass of about 150 kDa
(Fig. 2A). No band was detected
in the control (Fig. 2B), in
which the membrane was incubated with NB-PBS in place of the antibody.
|
Immunocytochemistry for Na+/K+-ATPase and
CFTR
A large number of Na+/K+-ATPase immunoreactive
chloride cells were detected in both the whole-mount preparations and
cryosections of the gill filaments in all experimental fish
(Fig. 3).
Fig. 1B shows the changes in
the average size of chloride cells following transfer. The chloride cells
became significantly larger at 12 h, but were decreased on day 7 to a size
equivalent to that before transfer. The decreased chloride cell size on day 7
was significantly increased again on day 14, and the increased cell size was
maintained thereafter.
|
CFTR immunoreactivity was detected in the apical region of Na+/K+-ATPase-immunoreactive chloride cells in seawater-acclimated fish before transfer in both whole-mount preparations and cryosections (Fig. 3A,B). The immunoreactivity appeared along the apical membrane, which formed a pit in seawater, and was rarely observed in the other part of the cell (Fig. 3A,B). The apically located CFTR immunoreactivity was also present in fish at 3 h after transfer to freshwater (Fig. 3C,D), but the intensity was apparently reduced at 12 h (Fig. 3E,F). The signal completely disappeared on day 1 and was not observed thereafter (Fig. 3G-L).
Scanning electron-microscopic observations
Chloride cells are in contact with the external environment through their
apical surface. The apical membranes of chloride cells were located at the
boundary of pavement cells, and most frequently observed on the afferent edge
of gill filament epithelia. In seawater-acclimated fish before freshwater
transfer, the apical membrane of most chloride cells invaginated to form a pit
(Fig. 4A,B). At 3 h after
transfer, the apical pits of chloride cells became shallow and the openings
were enlarged to a varying extent, whereas the apical structure of some
chloride cells was similar to that observed at 0
h(Fig. 4C,D). Through the
enlarged apical opening, poorly developed microvilli were observed on the
apical membrane, and a small apical surface of an accessory cell occasionally
appeared next to the chloride cells (Fig.
4D). At 12 and 24 h (day 1), the pit structures typically seen at
0 h were rarely observed, but the apical membrane of most chloride cells was
flat and equipped with microvilli (Fig.
4E-G). On day 3, however, some shallow pits with an enlarged
opening appeared again among the flat apical membranes of chloride cells
(Fig. 4H,I). On day 7 and
later, the apical membranes of chloride cells were flat or even protrusive,
and the microvilli on the surface were well developed
(Fig. 4J-M).
|
Transmission electron-microscopic observations
Chloride cells are generally characterized by a rich population of
mitochondria in the cytoplasm, and thus readily identified by TEM
(Fig. 5). In
seawater-acclimated fish, the apical membrane invaginated to form a pit, as
seen with SEM. The chloride cells often interdigitated with neighboring
accessory cells, forming multicellular complexes. The chloride and accessory
cells shared an apical pit (Fig.
5A). At 3 h after transfer, the pit became shallow and in some
cases, the apical region of chloride cells rose to the external environment.
The chloride cell and neighboring accessory cell still formed a cellular
complex (Fig. 5B). At 12 h, the
pit structures of chloride cells were rarely observed, and the apical
membranes were flat or slightly projecting. At the same time, accessory cells
were no longer observed at 12 h (Fig.
5C). The microvilli on the apical membrane were increasingly
developed after 12 h (Fig.
5D-H), and the apical membrane was covered with dense and
elongated microvilli on day 30 (Fig.
5H). As seen with SEM, exceptionally some chloride cells with a
shallow pit and an enlarged apical opening appeared on day 3, but were not
observed afterward. Moreover, numerous small electro-dense vesicles were
evident below the apical membrane in the cytoplasm of chloride cells on days
7-30 (Fig. 5F-H).
|
Chloride cell replacement following transfer from seawater to
freshwater
Many Rhodamine 123-positive chloride cells were detected in the
afferent-vascular edge of gill filaments in both seawater- and
freshwater-acclimated fish at the beginning of the experiment (day 0). After
the second staining with MitoTracker on day 3, Rhodamine 123-positive
(pre-existing) chloride cells and MitoTracker-positive (pre-existing and newly
differentiated) chloride cells were observed under a fluorescence microscope
as green and red cells, respectively (Fig.
6A,B,D,E,G,H).
|
There were no significant differences in number between Rhodamine 123-positive cells on day 0 and MitoTracker-positive cells on day 3 in the three experimental groups (Fig. 6C,F,I). In other words, the total number of chloride cells did not change in any experimental condition. Following transfer from seawater to freshwater, however, the number of Rhodamine 123-positive (pre-existing) cells decreased significantly (P<0.01) on day 3 (Fig. 6C), whereas there was no difference in the number of pre-existing cells between days 0 and 3 in the freshwater- or seawater-maintained groups (Fig. 6F,I). On day 3, the ratios of newly differentiated (Rhodamine-negative, MitoTracker-positive) chloride cells were 14.7%, 1.2% and 1.8% in transferred, seawater and freshwater groups, respectively. Significant differences in frequency of newly differentiated chloride cells were detected between freshwater-transferred and the two control groups (P<0.01), but not between seawater and freshwater control groups (Fig. 6C,F,I).
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Discussion |
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Chloride cells were detected in the whole-mount preparations of the gill
filaments by LSM. The mean size of chloride cells became significantly larger
just after transfer, when plasma Na+ levels showed a transient
decrease. The increase in the chloride cell size might be caused by the sudden
decrease in plasma ion concentration following direct transfer to the
hyposmotic environment. Most probably, enlarged chloride cells in freshwater
are in charge of ion uptake to compensate for the ion loss. Enlargement of
chloride cells has been reported in both gills and opercular membrane of
freshwater-adapted killifish (Katoh et
al., 2001), whereas chloride cells are larger in seawater than in
freshwater in other euryhaline species
(Langdon and Thorpe, 1985
;
Richman et al., 1987
; Uchida
et al., 1996
,
2000
;
Shiraishi et al., 1997
;
Hiroi et al., 1999
). In
killifish gills, seawater-type chloride cells have an apical pit and
interdigitate with neighboring accessory cells to form multicellular
complexes, whereas the apical membrane is flat or protrusive with developed
microvilli on it in freshwater-type cells
(Katoh et al., 2001
). A
similar apical membrane structure has been observed in several species adapted
to freshwater (Hossler et al.,
1985
; Laurent and Hebibi,
1988
; Perry et al.,
1992
; Shawn et al.,
1993
; Greco et al.,
1996
; Perry, 1997
,
1998
;
Kelly et al., 1999
).
In spite of the occurrence of distinct freshwater- and seawater-type chloride cells, whether one cell type degenerates and is replaced by newly differentiated cells of another type, or whether chloride cells alter their functions in response to environmental salinity change, is still controversial. In addition to typical seawater- and freshwater-type cells, our SEM and TEM observations revealed the occurrence the intermediate type, which was most frequently observed at 3 h after transfer. The intermediate-type cells were accompanied by accessory cells, which is characteristic of the seawater type. However, the apical openings were larger and the pits were shallower than those observed in the typical seawater type, and more like those in the freshwater type. Through the enlarged opening, poorly developed microvilli were observed on the apical membrane, which was not evident in the seawater type. Considering that the intermediate-type cells appeared within 3 h after the transfer into freshwater, it is most probable that seawater-type cells were transformed into the intermediate type. Since freshwater-type cells, similar to those in fish fully acclimated to freshwater, were frequently observed at 12 h, the intermediate-type cells appear to be further transformed into the freshwater type. Thus, our observations indicate a plasticity of chloride cells, whereby in killifish the seawater type is transformed into the freshwater type following transfer to freshwater.
It has been demonstrated in some teleosts that chloride cells alternate
their morphology and ion-transporting functions to meet abrupt environmental
osmotic changes. Using a sequential observation technique, Hiroi et al.
(1999) revealed that single
freshwater-type chloride cells in the yolk-sac membrane of Mozambique tilapia
Oreochromis mossambicus are transformed into seawater-type
multicellular complexes during seawater adaptation. Mudskipper
Periophthalmus modestus tolerates hypotonicity by closing the apical
pit of existing seawater-type chloride cells on exposure to freshwater
(Sakamoto et al., 2000
). The
apical pit density in the opercular membrane decreases in seawater-adapted
killifish subjected to a hyposmotic shock on the basolateral surface
(Daborn et al., 2001
). All
these findings are interpreted as an acute adaptive response of chloride cells
to environmental changes.
The CFTR has been identified electrophysiologically in the apical membranes
of killifish chloride cells (Marshall et
al., 1995), and has been cloned and sequenced from the gills of
seawater-adapted killifish (Singer et al.,
1998
). According to the sequence analysis, killifish CFTR has the
same carboxy-terminal sequence as human CFTR
(Singer et al., 1998
), and
thus a monoclonal antibody to human CFTR directed against this epitope is
applicable to killifish. In our western blot analysis, the antibody recognized
one specific protein band, of approx. 150 kDa, in agreement with the expected
size of killifish CFTR (Singer et al.,
1998
). The result therefore confirmed the high specificity and
availability of the antibody to killifish CFTR.
Marshall et al. (1999)
reported that killifish CFTR expression increases at 8 h after transfer from
freshwater to seawater, peaking at 24 h and remaining at levels higher than
those in freshwater after 30 days in seawater. The CFTR expression was linked
to the enhancement of Cl- secretion that occurred at 24 h after
transfer to seawater. In the present study, the CFTR immunoreactivity was
detected in the apical membrane of chloride cells at 0, 3 and 12 h after
transfer from seawater to freshwater, but disappeared at 24 h. This result is
consistent with the observation by Marshall et al.
(1999
), in the sense that CFTR
expression in chloride cells is more closely related to seawater adaptation,
suggesting the involvement of CFTR-immunoreactive chloride cells in
Cl- secretion in seawater. This also indicates that pre-existing
seawater-type chloride cells are able to change their function, as well as
their morphology, to those of the freshwater-type.
The recovery of plasma Na+ levels from a sharp decrease just
after transfer to freshwater could be partly accounted for by the functional
and morphological alteration of chloride cells from seawater to freshwater
type. In addition to such an acute response, we examined chloride cell
replacement as a long-term effect of freshwater transfer. In the present
study, we developed the time-differential double fluorescent staining (TDS)
technique, which made it possible to visualize chloride cell replacement more
directly and easily than by conventional methods. Fluorescent labeling
techniques provide a fast and valuable method for qualitatively and
quantitatively assessing chloride cells, and mitochondria-specific probes
enabled the vital staining of chloride cells
(Ayson et al., 1994;
Hiroi et al., 1999
;
Li et al., 1995
;
Sakamoto et al., 2000
;
Marshall et al., 2002
). For
the TDS, we adopted two specific mitochondrial probes, Rhodamine 123 and
MitoTracker (MitoTracker Red CM-H2XRos), to track the
turnover of branchial chloride cells.
Although the total number of chloride cells did not change in any group during the experimental 3 day period, the ratio of newly differentiated chloride cells to the total cells at the end of the experiment was markedly higher in the freshwater-transferred group (14.7%) than the freshwater- or seawater-maintained group (1.8% and 1.2%, respectively). Meanwhile, there was no significant difference between numbers in seawater- and freshwater-maintained groups. These results indicate that the turnover rate of chloride cells is relatively low under a constant osmotic environment, but is accelerated after transfer to freshwater. Under constant osmotic conditions, a small proportion of chloride cells are continuously replaced with the same type of cells. In contrast, the enhanced chloride cell turnover after transfer seems to be the result of replacement of pre-existing seawater-type chloride cells with newly differentiated, freshwater-type cells. Thus, the replacement of chloride cells after transfer not only sustains the chloride cell population, but also contributes to the alteration in ion-transporting function of the gills. This may serve as a long-term adaptative response to a different osmotic environment, together with a short-term response of morphological and functional transformation of pre-existing chloride cells. During the time-course observations on chloride cells after transfer into freshwater, a rapid increase in the chloride cell size at 12 and 24 h was followed by a gradual decrease over 1 to 3 days after transfer. In our SEM and TEM observations, developing freshwater-type cells were occasionally observed on day 3. Such transient inactivation of chloride cells can be explained by the recruitment of newly differentiated, small freshwater-type cells, following the short-term adaptive response.
Our results contrast sharply with those from some other teleosts where the
chloride cell turnover was not different between freshwater and seawater
killifish. In guppy, cell differentiation and renewal of the gill epithelium
were three times faster in 50% seawater-adapted fish than in
freshwater-adapted fish (Chretien and
Pisam, 1986). Using 5-bromo-2'-deoxyuridine (BrdU)
incorporated into nuclei during DNA synthesis, Uchida and Kaneko
(1996
) revealed that the
chloride cell turnover of chum salmon was about three times greater in
seawater than in freshwater. This discrepancy between killifish and other
fishes may reflect the preference of killifish for seawater over freshwater.
The Na+/K+-ATPase activity and oxygen consumption are
generally greater in seawater than in freshwater in most euryhaline teleosts,
such as tilapia, salmonids and eels
(Morgan et al., 1997
; Uchida
et al., 1996
,
2000
;
Richman et al., 1987
;
Sasai et al., 1998
),
suggesting their freshwater preference. In these species, the chloride cell
turnover seems greater in seawater, in which chloride cells consume more
energy for adaptation to the less favorable environment. In contrast, there
was no difference in either Na+/K+-ATPase activity or
oxygen consumption between freshwater- and seawater-adapted killifish
(Katoh et al., 2001
). Since
chloride cells are equally active in both media, the turnover rates may not be
different between freshwater- and seawater-acclimated killifish.
In this study, we focused on the short- and long-term adaptive responses of killifish following direct transfer from seawater to freshwater. Just after transfer, when plasma Na+ levels showed an abrupt decrease, the transformation of chloride cells occurred from the pre-existing seawater type to freshwater type via the intermediate type. This transformation process is accompanied by the disappearance of apically located Cl- channel (CFTR) and neighboring accessory cells. Such morphological and functional changes are interpreted as an acute adaptive response of chloride cells to cope with the unexpected decrease in plasma ion levels following direct transfer to freshwater. On the other hand, the chloride cell turnover was enhanced after transfer, and pre-existing seawater-type chloride cells are replaced with newly differentiated, freshwater-type cells. This is considered a long-term adaptive response to the hyposmotic environment. Taken together, these results clearly indicate that the branchial chloride cells show a two-phase response in adapting to freshwater environment: the short-term functional and morphological transformation of pre-existing cells, and the long-term cell replacement by newly differentiated cells. Moreover, for examination of the chloride cell replacement, we have established the TDS technique. Compared with conventional methods using 3H-thymidine or BrdU, the newly developed TDS technique provides a more convenient and precise method for assessing chloride cell turnover. Since limited information is available on chloride cell turnover, this new technique will be an excellent experimental tool for further studies on chloride cell differentiation.
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