Mitochondria-rich cell activity in the yolk-sac membrane of tilapia (Oreochromis mossambicus) larvae acclimatized to different ambient chloride levels
1 Graduate Institute of Life Sciences, National Defense Medical Center,
Nei-Hu, Taipei 114, Taiwan, ROC
2 Institute of Zoology, Academia Sinica, Nankang, Taipei 11529, Taiwan,
ROC
* Author for correspondence (e-mail: pphwang{at}gate.sinica.edu.tw)
Accepted 6 January 2004
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: mitochondria-rich cell, MRC, yolk sac, tilapia, Oreochromis mossambicus, larva, ambient chloride
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
MRC morphology and function in freshwater teleosts have previously been
studied in depth, and several studies have focused on the apical membrane
(often forming an apical crypt) configuration of MRCs in response to external
or internal changes (Perry,
1997; Van Der Heijden et al.,
1997
,
1999
;
Marshall, 2002
;
Wilson and Laurent, 2002
).
Acidification of water increases the fractional surface area of gill MRCs
without changing MRC density, indicating morphological responses of the
epithelium to environment changes (Laurent
and Perry, 1991
), and this could result from responses of either
the MRCs, a rearrangement of the pavement cells or a combination of these
phenomena. Metabolic alkalosis (bicarbonate infusion) and cortisol treatment
increase the number and fractional surface area of exposed MRCs in freshwater
rainbow trout (Oncorhynchus mykiss;
Perry and Goss, 1994
). Laurent
et al. (1995
) examined the
Lake Magadi tilapia (Oreochromis alkalicus grahami), a species
uniquely adapted to severely alkaline freshwater, which possesses
well-developed gill MRCs with apical crypts that remain open in alkaline water
(pH 10) but close in neutral water (pH 7; within 23 h of exposure).
Crypts appear again upon long-term (24 h) residence in neutral water, and this
demonstrates the dynamics of MRC apical membrane exposure.
It is well known that the apical membrane of MRCs forms an apical crypt
configuration in seawater fishes but may be flush with or raised slightly
above adjacent pavement cells in most freshwater-acclimated fishes
(Perry and Laurent, 1993);
however, crypts have been reported for freshwater Mozambique tilapia
(Oreochromis mossambicus) as well
(Lee et al., 1996
;
Van der Heijden et al., 1997
)
and this may illustrate our limited understanding of MRC biology.
In a previous study on freshwater Mozambique tilapia, we categorized
subtypes of MRCs with different apical surfaces as wavy-convex, shallow-basin
and deep-hole (crypts) and correlated subtype abundance with medium
Ca2+, Na+ and Cl concentrations
(Lee et al., 1996;
Chang et al., 2001
).
These subtypes of MRCs were suggested to be functionally different for ion
transport.
More recently, we found that the morphology of these apical surfaces
changed in a matter of hours with variations in ambient Cl
concentrations and correlated with the rate of Cl influx in
tilapia, which suggests that MRC morphology reflects distinct capabilities for
Cl uptake (Lin and
Hwang, 2001; Chang et al.,
2003
). In the present study, we provide further cytological
evidence for the hypothesis. When ambient chloride levels are extremely
reduced, gradually more active MRCs are observed and the cells enlarge their
surface area, probably to upregulate their Cl uptake
capacity; conversely, when ambient Cl levels are increased,
MRCs are inactivated by constriction of their apical openings and become
totally covered by adjacent apical pavement cells. We used immunocytochemistry
and vital staining to trace the turnover of yolk-sac MRCs in tilapia larvae
acclimated to high or low ambient Cl levels. We traced MRCs
by Na+/K+-ATPase immunostaining and scored them as
active (in contact with the water; Van der Heijden et al.,
1997
,
1999
) when they could be
labeled with Con-ATexas-Red; we then examined the changes in active
cells during chloride acclimation. In addition, yolk-sac MRCs were traced with
DASPEI vital staining to examine turnover and interaction with adjacent
pavement cells.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Three artificial freshwater media were made (1) control, reflecting local tap water, (2) with low Cl content and (3) with high Cl content by addition of appropriate amounts of NaCl, Na2SO4, MgSO4, K2HPO4, KH2PO4 and CaSO4 to deionized water (see Table 1). The temperature of the media was kept between 26°C and 28°C. During experiments, larvae were not fed; media were changed daily to guarantee optimal water quality.
|
Immunohistochemistry
Larvae were anaesthetized on ice and fixed in 4% paraformaldehyde in 0.1
mol l1 phosphate buffer (PB; pH 7.4) for 30 min at 4°C.
After washing with phosphate-buffered saline (PBS; pH 7.4), fixed larvae were
incubated in 1 mg ml1 Texas-Red-conjugated concanavalin-A
(Con-A; Molecular Probes, Eugene, OR, USA; dissolved in redistilled water) for
30 min at room temperature (RT; 2427°C). After rinsing for 20 min
at 4°C, the larvae were further fixed and permeabilized with 70% ethanol
at 20°C for 10 min. After washing with PBS, samples were incubated
overnight at 4°C with a monoclonal antibody against the
5-subunit of the chicken Na+/K+-ATPase
(Developmental Studies Hybridoma Bank, University of Iowa) diluted 1:200 with
PBS containing 10% normal goat serum and 1% bovine serum albumin. After
rinsing with PBS for 20 min, the larvae were further incubated in goat
anti-mouse IgG conjugated with fluorescein isothiocyanate (FITC; Jackson
Immunoresearch Laboratories, West Grove, PA, USA; dilution 1:100) for 2 h at
RT. After staining, the whole larvae were mounted in an observation chamber
that was composed of a cover slip (24 mmx24 mm) and spacers slightly
thinner than the thickness of the yolk sac of larvae. In this situation, the
slightly compressed yolk sac provided a flatter area for observation.
Observations and image acquisitions were made using a Leica TCS-NT confocal
laser scanning microscope (Leica Lasertechnik, Heidelberg, Germany) equipped
with 10x/0.3, 20x/0.4, 40x/1.2 and 100x/1.35
(magnification/numerical aperture) objective lenses and appropriate filter
sets for simultaneous monitoring of FITC and Texas Red. Cross-talk between the
two fluorescent signals was negligible in our system.
Determination of active MRCs
Two-day-old larvae were transferred from normal freshwater to the three
artificial freshwater media. After 0 h (normal freshwater controls), 24 h or
48 h, larvae were sampled to determine active MRCs. MRCs were double-labeled
with antibody against the sodium pump subunit and
Con-ATexas-Red as described above. Confocal laser scanning was
performed on the yolk-sac area of whole-mounts of larvae. Three areas (0.25
mm2) per individual were scanned with a 20x/0.4 objective
lens, and the acquired images were further enhanced and analyzed with
MetaMorph software (Universal Imaging Corporation, Philadelphia, PA, USA). The
densities of MRCs and their exposed apical surfaces were separately
quantified, and the ratio of active MRCs over total MRCs was calculated as
follows: active MRC (%) = (density of exposed apical surfaces/density of
MRC)x100%.
Vital staining and time-sequential tracing of MRCs
Two-day-old larvae were incubated in normal freshwater containing 300
µmol l1 2-(4-dimethylaminostyryl)-1-methylpyridinium
iodide (DASPEI; Sigma, St Louis, MO, USA) for 2 h at 27°C. After
incubation, larvae were rinsed with normal freshwater and then anesthetized
with MS-222 (3-aminobenzoic acid ethyl ester; Sigma; 0.1 mg
l1; pH 7.0, buffered with 3 mmol l1 MOPS)
for 5 min prior to observation by confocal microscopy. An individual larva was
mounted in an observation chamber that was composed of a cover slip (24
mmx24 mm) and spacers (made with stacking cover slips) slightly thinner
than the thickness of the yolk sac of the larvae. This chamber is similar to
that used in a previous report by Hiroi et al.
(1999). The flattened yolk-sac
surface of the mounted larva was then analyzed by confocal laser scanning
microscopy. One area (0.25 mm2) per individual was selected for
image acquisition. Usually one or more melanophores in the center of the area
served as a marker to recognize the area under study during subsequent
analyses. After scanning, the larvae were removed from the chamber and placed
in one of several artificial freshwater tanks for further incubation. The
scanning procedure was usually finished within 5 min to keep damage to the
larvae to a minimum. Nine individuals from the same brood were examined in
this experiment. After initial scanning, these larvae were transferred to the
three media (three larvae per medium) for further incubation. After 12 h and
24 h ofincubation, the same areas of the larvae were scanned again, and
DASPEI-positive cells were scored. Two hours prior to the second and third
scanning, DASPEI (300 µmol l1) was added to the
incubation media. During the experiments, neither mortality nor significant
damage occurred in the traced larvae.
Apical membrane internalization in MRCs
Live, 2-day-old larvae were immersed in a Con-A solution (1 mg
Con-ATexas-Red dissolved in 1 ml artificial water) for 30 min. Then,
the larvae were transferred to Con-A-free artificial water and gently washed
using a wide-mouth pipette for 3 min. After washing, the larvae were kept in
artificial water, shielded from light for 2 h. Then, the larvae were
sacrificed and immunolabeled for Na+/K+-ATPase as
described above.
Simultaneous labeling of MRCs, MRC apical membranes and pavement cells
Live, 2-day-old larvae were stained with Con-A in vivo as
described above. After washing, larvae were transferred to water with a high
Cl content for 24 h, shielded from light to avoid
photobleaching of the fluorescent probes used. After 24 h, 300 µmol
l1 DASPEI was added to the medium for 30 min to stain the MR
cells. After washing, the larvae were transferred to an FM1-43
{N-(3-triethylammoniumpropyl)-4-[4-(dibutylamino)styryl]pyridinium
dibromide; Molecular Probes} solution (5 µmol l1; diluted
in the same medium) for 20 min to stain pavement cells.
After staining, larvae were anesthetized with MS-222 and immediately scanned with a confocal microscope. In pilots, larvae had been stained with various concentrations of FM1-43 over increasing times, which yielded similar staining intensities but increasing fluorescence intensities with concentration and time. Treatment with 5 µmol l1 FM1-43 for 20 min was selected as the optimal condition for our confocal analyses.
Statistics
Values were compared using one-way analysis of variance (ANOVA) by Tukey's
pair-wise method. Values are presented as means ± standard deviation
(S.D.).
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Active and inactive MR cells
When larvae were double-labeled with Na+/K+-ATPase
antibody and Con-ATexas-Red, we noticed that a portion of MRCs (rich in
Na+/K+-ATPase labeling) were not labeled by Con-A and
thus concluded that such cells are not in contact with the water
(Fig. 2A). In previous studies
(Lin and Hwang, 2001;
Chang et al., 2003
), increased
density of MRCs and increased exposed surface area of the apical membrane of
MRCs were shown to correlate closely and positively with Cl
influx. Based on these observations, Con-A-negative MRCs are referred to here
as `inactive' cells, in contrast to Con-A-positive, `active' MRCs. The
proportion of active MRCs of the total number of MRCs
(Fig. 2) was calculated and
compared between different groups of larvae acclimatized to the three media.
Only
20% of yolk-sac MRCs in 2-day-old control larvae are active, and
this proportion increased slightly but significantly to 24% with development
from 2 to 4 days old (Fig. 3A). However, in larvae acclimated to low Cl water, active MRCs
increased dramatically from 20% to 45% during a 48 h acclimatization. By
contrast, active MR cells declined to
13% after 48 h in high
Cl water (Fig.
3A). Thus, external Cl levels can alter the
proportion of active MRCs in the yolk-sac membrane. Indeed, the density of
total MRCs (active and inactive cells) in the yolk-sac membrane was similar
among the three groups of larvae (Fig.
3B).
|
|
MRC dynamics
MRCs in yolk-sac membrane vitally stained with DASPEI were sequentially
scanned with a confocal microscope at 12-h intervals for 24 h. During
development, the yolk sacs of larvae are, of course, gradually absorbed and,
during absorption, the MRCs in the yolk sac are found to a greater extent in
the vicinity of the larval body trunk. However, individual MRCs keep their
position relative to one another during yolk sac absorption and could be
easily identified at the 12-h intervals of scanning.
Fig. 4 shows confocal images of
sequentially (0 h, 12 h and 24 h) scanned MRCs in larvae transferred from
normal to high-Cl water. More than 90% of labeled cells
survived the acclimation and scanning procedures (the white-numbered cells in
the images; Figs 4,
5). About 10% of the cells
observed over the 24 h period were new, small DASPEI-positive cells
(yellow-numbered cells, Figs 4,
5). Interestingly, no
significant change in DASPEI-positive cell numbers was observed in the other
two groups of larvae (normal and low Cl water), an
indication that cell turnover was not affected by a decrease in ambient
Cl levels (Fig.
5).
|
|
Internalization of the apical membrane in MRCs
As shown by confocal optical sectioning in
Fig. 6, in larvae pre-stained
with Con-A and then incubated in normal water for 2 h, the Con-A-labeled
apical membrane is internalized in MRCs, evidenced by punctate Con-A label
(vesicles) beneath the apical membrane of MR cells
(Fig. 6A).
Fig. 6B,C shows optical
sections 3 µm and 6 µm beneath the apical surface.
|
FM1-43 staining: pavement cells and apical exposure of MRCs
The lipophilic vital stain FM1-43 was used as a membrane marker to label
the cell membrane of the integument epithelium. In larvae exposed to the dye
in water, FM1-43 labeled mainly the integument (pavement) cells of the
yolk-sac membrane, revealed as a layer of flat polygonal cells with very
clearly stained cell boundaries (Fig.
7A). At high magnification (1000x), these squamous,
polygonal cells showed typical characteristics of pavement cells with
microridges; these cells form the outermost cell layer of the epithelium and
cover the MRCs except for their apical surfaces
(Fig. 7B). FM1-43 penetrates
the pavement cell membrane and labels organelles inside pavement cells. When
adjusting the focal plane to 35 µm beneath the surface of pavement
cells, we observed strong signals of organelles surrounding the nuclei of
pavement cells (Fig. 7C). With
this staining technique, proliferating pavement cells (appearing to have
23 nuclei) were occasionally observed in yolk-sac membranes
(Fig. 7C).
|
As the Con-A-labeled apical membrane of MRCs become internalized, we then set out to transfer larvae pulse-labeled with Con-A from normal water to high Cl water for 24 h. After this 24-h period, still-internalized Con-A vesicles were seen in MRCs, as confirmed first by DASPEI labeling. Next, larvae were stained with FM1-43 to examine the behavior of the MRCs and the pavement cells during the acclimation (Fig. 7DF). Confocal optical sections through the surface plane (pavement cells; Fig. 7D), through a plane 6 µm beneath the surface (MRCs; Fig. 7E) and the merged picture of the two sections (Fig. 7F) showed MRCs containing Con-A-labeled vesicles (indicating that these cells had previously been in contact with the water) fully covered by pavement cells. Apparently, a high Cl medium forms a stimulus for active MRCs to withdraw from the surface and avoid contact with the water. In addition, the above data infer that pavement cells adjacent to MRCs fill the gaps when these cells withdraw and become inactive (no more direct contact with the ambient medium).
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We assumed that the Con-A-labeled MRCs were `active' MRCs because they were
in contact with the water through the apical surfaces, which were shown to
correlate with the activity of Cl uptake in tilapia
(Lin and Hwang, 2001;
Chang et al., 2003
).
Conversely, MRCs completely covered by pavement cells were presumed to be
`inactive' in Cl uptake. In addition to the correlation
between the MRC morphology and the whole-body
36Cl uptake
(Lin and Hwang, 2001
), our
recent work using a scanning ion-selective electrode technique to probe
Cl flux in yolk-sac MRCs of tilapia has demonstrated that
these unexposed MRCs are functionally inactive (L. Y. Lin and P. P. Hwang,
unpublished data).
Several previous studies have employed fluorochrome-conjugated lectin to
label the apical surfaces of MRCs. Li et al.
(1995), Van Der Heijden et al.
(1997
) and Lee et al.
(2000
) used
fluorochrome-conjugated Con-A, which binds specifically to
-glucopyranosyl glycoprotein residues, to localize the exposed apical
surfaces of MRCs identified with either mitochondrial staining or an
Na+/K+-ATPase marker. Wheat germ agglutinin (WGA), which
specifically binds to N-acetylglucosamine and
N-acetylneuraminic acid residues, was also used by Tsai and Hwang
(1998
) to identify specific
subpopulations of MRCs. Recently, Goss et al.
(2001
) used another lectin,
peanut agglutinin (PNA; which binds specifically to terminal ß-galactose
residues), to separate two subtypes of MRCs. Con-ATexas-Red has been
shown to bind strongly to apical surfaces (also termed apical crypts in
previous reports) of tilapia MRCs and has been used as a marker to identify
exposed or mature MRCs (Li et al.,
1995
; Van Der Heijden et al.,
1997
; Lee et al.,
2000
). In the present study, Con-ATexas-Red was also shown
to bind specifically to apical surfaces of yolk-sac MRCs in tilapia larvae
(Fig. 1). We could easily
identify wavy-convex MRCs, particularly in low-Cl larvae.
However, it was more difficult to discriminate the shallow-basin and deep-hole
types due to the similar staining shape. By using Con-ATexas-Red and
Na+/K+-ATPase antibody double-labeling, we scored the
active MRCs and examined their profiles in larvae acclimated to high- or
low-Cl water. In normal larvae 24 daysafter hatching,
2024% of MR cells were Con-A positive (active), which is comparable
with previous data determined by counting MRCs in contact with the water
environment on tissue sections of tilapia larvae
(Van Der Heijden et al.,
1999
). The proportion of active MRCs increased from 20% to 45%
after 48-h low-Cl acclimation but declined from 20% to 13%
after high Cl acclimation. The changing profiles are
congruent with our previous counting of exposed MRCs with scanning electronic
microscopy (SEM; Lin and Hwang,
2001
). However, using this double-labeling technique, we could
score the active and inactive MRCs simultaneously.
In the present study, we labeled MRCs with DASPEI vital stain and
sequentially monitored their turnover in intact animals during acclimation to
both high- and low-Cl water. This method was modified from a
previous report by Hiroi et al.
(1999), who used it to
investigate MRC turnover in tilapia larvae during seawater acclimation and
found that a portion of freshwater-type MRCs are able to transform to
seawater-type MRCs. The vital stain DASPEI and its analogue DASPMI, which
accumulate in active mitochondria, are the most common dyes used to label MRCs
(Li et al., 1995
;
Witters et al., 1996
;
Rombough, 1999
;
Van Der Heijden et al., 1999
).
Using this technique here, we found that MRC turnover was not altered by
ambient chloride changes, suggesting that the change in the densities and
subtypes of MRCs induced by ambient chloride
(Lin and Hwang, 2001
) reflects
the process of MRCs undergoing structural and functional modification.
FM1-43, which was originally developed as a membrane potential sensor, has
become an increasingly useful tool in the study of membrane trafficking,
synaptic vesicle recycling and synaptic transmission
(Cochilla et al., 1999). In
addition, when used as a marker of cell membranes and surface areas, FM1-43 is
useful for determining the means by which damaged cells are able to repair
their membranes. Studies on endothelial cells
(Miyake and McNeil, 1995
), sea
urchin eggs and embryos (Steinhardt et
al., 1994
) and crayfish medial giant axons
(Eddleman et al., 1997
)
indicate that at the site of an injury, exocytosis of lipid vesicles is used
to repair holes in the cellular membrane. In the present study, we used FM1-43
as a membrane marker to label the apical membrane of the epithelium covering
the larval skin. We found that FM1-43 effectively stained the apical
microstructure of pavement cells (microridges) and clearly outlined the
boundary of these cells. Interestingly, we found that FM1-43 seemed to
permeate the apical membrane of pavement cells and consequently stained the
membrane organelles inside these cells. Since FM1-43 is used for tracing
membrane trafficking (exocytosis and endocytosis), we also checked whether the
vesicles inside these cells are internalized from the labeled apical membrane
through endocytosis. It seems unlikely that the considerable number of
vesicles is internalized from the apical membrane in such a short time
(
20 min); in addition, no obvious vesicle trafficking was observed inside
these cells. However, FM1-43 did not permeate MR cells but only slightly
stained their apical membranes. Applying the same method to zebrafish
(Danio rerio) larvae, we found that FM1-43 could not penetrate the
apical membrane of pavement cells, implying that the staining property is cell
type and/or species dependent. A recent report found that FM1-43 also
penetrates specific types of sensor neurons through mechanotransduction
channels (Meyers et al.,
2003
). Whether or not a similar channel or conducting pathway for
FM1-43 exists in tilapia pavement cells needs to be further investigated.
With FM1-43 vital staining, we could easily identify the apical surfaces of MRCs surrounded by adjacent pavement cells, similar to what we observed with SEM. However, this method allows us to investigate the morphology of pavement cells and apical surfaces of MRCs in live animals, which cannot be achieved with SEM. Using FM1-43 and DASPEI double-staining, we could simultaneously trace these two types of cells (pavement cells and MR cells) in a live animal and examine their interaction. We found that high-Cl water stimulated active MRCs to withdraw from the surface and avoid contact with the water. Pavement cells adjacent to MRCs play the role of filling the gaps when MRCs withdraw and become inactive.
A similar regulatory mechanism has been proposed by Goss and colleagues
(Goss et al., 1992,
1998
;
Goss and Perry, 1993
;
Perry and Goss, 1994
) in their
studies of freshwater MRCs responding to respiratory acidbase
disturbances. The present work supports their model and has adapted this
regulatory mechanism to freshwater tilapia responding to ambient
Cl disturbances. Recently, Daborn et al.
(2001
) used opercular membranes
of killifish (Fundulus heteroclitus) to examine the interactions
between pavement cells and MR cells during abrupt salinity changes and also
concluded that osmotic shock caused MRCs to adjust their apical surface size
by interacting with adjacent pavement cells. The actin cytoskeleton of MRCs
was shown to maintain the apical surfaces required for modification of the
surfaces in response to osmotic shock
(Daborn et al., 2001
). In the
present study, internalization of the apical membrane of Con-A-labeled MR
cells was observed, indicating that apical membrane turnover might also be
involved in maintaining and regulating the structure and composition of apical
surfaces. Numerous vesicles observed beneath the apical membrane of MRCs in
previous reports investigating the subcellular structure of MRCs might imply
the active turnover of apical membranes. The physiological significance of
apical membrane turnover requires further investigation.
It has been generally accepted that MRCs that are not exposed to the
external environment are undergoing differentiation and they have been
considered to be functionally `immature' MRCs (Van Der Heijden et al.,
1997,
1999
). However, in our
findings and other reports (Laurent et
al., 1995
; Sakamoto et al.,
2000
; Daborn et al.,
2001
), these MRCs could be inactivated from functional cells due
to being covered by pavement cells responding to ambient or internal
ion/osmolarity alterations. Therefore, we use `inactive' to describe these
unexposed MRCs, which may be composed of immature and inactivated MRCs. Once
upregulation of ion uptake is required, both of these forms might be recruited
into becoming functionally active MR cells.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Ayson, F. G., Kaneko, T., Hasegawa, S. and Hirano, T. (1994). Differential expression of two prolactin and growth hormone genes during early development of tilapia (Oreochromis mossambicus) in fresh water and seawater: implications for possible involvement in osmoregulation during early life stages. Gen. Comp. Endocrinol. 95,143 -152.[CrossRef][Medline]
Chang, I. C., Lee, T. H., Yang, C. H., Wei, Y. Y., Chou, F. I. and Hwang, P. P. (2001). Morphology and function of gill mitochondria-rich cells in fish acclimated to different environments. Physiol. Biochem. Zool. 74,111 -119.[CrossRef][Medline]
Chang, I. C., Wei, Y. Y., Chou, F. I. and Hwang, P. P. (2003). Stimulation of Cl uptake and morphological changes in gill mitochondria-rich cells in freshwater tilapia (Oreochromis mossambicus). Physiol. Biochem. Zool. 76,544 -552.[CrossRef][Medline]
Cochilla, A. J., Angleson, J. K. and Betz, W. J. (1999). Monitoring secretory membrane with FM1-43 fluorescence. Annu. Rev. Neurosci. 22,1 -10.[CrossRef][Medline]
Daborn. K., Cozzi, R. R. and Marshall, W. S.
(2001). Dynamics of pavement cellchloride cell
interactions during abrupt salinity change in Fundulus heteroclitus.J. Exp. Biol. 204,1889
-1899.
Eddleman, C. S., Ballinger, M. L., Smyers, M. E., Godell, C. M.,
Fishman, H. M. and Bittner, G. D. (1997). Repair of
plasmalemmal lesions by vesicles. Proc. Natl. Acad. Sci.
USA 94,4745
-4750.
Goss, G. G., Adamia, S. and Galvez, F. (2001).
Peanut lectin binds to a subpopulation of mitochondria-rich cells in the
rainbow trout gill epithelium. Am. J. Physiol. Regul. Integr. Comp.
Physiol. 281,R1718
-R1725.
Goss, G. G. and Perry, S. F. (1993). Physiological and morphological regulation of acid-base status during hypercapnia in rainbow trout (Oncorhynchus mykiss). Can. J. Zool. 71,1673 -1680.
Goss, G. G., Perry, S. F., Wood, C. M. and Laurent, P. (1992). Mechanisms of ion and acid-base regulation at the gills of freshwater fish. J. Exp. Zool. 263,143 -159.[Medline]
Goss, G. G., Perry, S. F., Fryer, J. N. and Laurent, P. (1998). Gill morphology and acid-base regulation in freshwater fishes. Comp. Biochem. Physiol. A 119,107 -115.
Guggino, W. B. (1980). Salt balance in embryos of Fundulus heteroclitus and F. bermudae adapted to seawater. Am. J. Physiol. 238,R42 -R49.[Medline]
Hiroi, J., Kaneko, T. and Tanaka, M. (1999). In
vivo sequential changes in chloride cell morphology in the yolk-sac membrane
of Mozambique tilapia (Oreochromis mossambicus) embryos and larvae
during seawater adaptation. J. Exp. Biol.
202,3485
-3495.
Hwang, P. P., Lee, T. H., Weng, C. F., Fang, M. J. and Cho, G. Y. (1999). Presence of Na-K-ATPase in mitochondria-rich cells in the yolk-sac epithelium of larvae of the teleost Oreochromis mossambicus. Physiol. Biochem. Zool. 72,138 -144.[CrossRef][Medline]
Hwang, P. P. and Sun, C. M. (1989). Putative role of adenohypophysis in the osmoregulation of tilapia larvae (Oreochromis mossambicus; Teleostei): an ultrastructure study. Gen. Comp. Endocrinol. 73,335 -341.[Medline]
Hwang, P. P., Tsai, Y. N. and Tung, Y. C. (1994). Calcium balance in embryos and larvae of the freshwater-adapted teleost, Oreochromis mossambicus. Fish Physiol. Biochem. 13,325 -333.
Laurent, P., Maina, J. N., Bergman, H. L., Narahara, A., Walsh, P. J. and Wood, C. M. (1995). Gill structure of a fish from an alkaline lake: effect of short-term exposure to neutral conditions. Can. J. Zool. 73,1170 -1181.
Laurent, P. and Perry, S. F. (1991). Environmental effects on gill morphology. Physiol. Zool. 64,4 -25.
Lee, T. H., Hwang, P. P., Lin, H. C. and Huang, F. L. (1996). Mitochondria-rich cells in the branchial epithelium of the teleost, Oreochromis mossambicus, acclimated to various hypotonic environments. Fish Physiol. Biochem. 15,513 -523.
Lee, T. H., Hwang, P. P., Shieh, Y. E. and Lin, C. H. (2000). The relationship between "deep hole" mitochondria-rich cells and salinity adaptation in the euryhaline teleost Oreochromis mossambicus. Fish Physiol. Biochem. 23,133 -140.[CrossRef]
Li, J., Eygensteyn, J., Lock, R. A. C., Verbost, P. M., Van Der Heijden, A. J. H., Wendelaar Bonga, S. E. and Flik, G. (1995). Branchial chloride cells in larvae and juveniles of freshwater tilapia Oreochromis mossambicus. J. Exp. Biol. 198,2177 -2184.[Medline]
Lin, L. Y. and Hwang, P. P. (2001). Modification of morphology and function of integument mitochondria-rich cells in tilapia larvae (Oreochromis mossambicus) acclimated to ambient chloride levels. Physiol. Biochem. Zool. 74,469 -476.[CrossRef][Medline]
Marshall, W. S. (2002). Na+, Cl, Ca2+ and Zn2+ transport by fish gills: retrospective review and prospective synthesis. J. Exp. Zool. 293,264 -283.[CrossRef][Medline]
Meyers, J. R., MacDonald, R. B., Duggan, A., Lenzi, D.,
Standaert, D. G., Corwin, J. T. and Corey, D. P.
(2003). Lighting up the senses: FM1-43 loading of sensory cells
through nonselective ion channels. J. Neurosci.
23,4054
-4065.
Miyake, K. and McNeil, P. L. (1995). Vesicle accumulation and exocytosis at sites of plasma membrane disruption. J. Cell Biol. 131,1137 -1145.
Perry, S. F. (1997). The chloride cell: structure and function in the gills of freshwater fishes. Annu. Rev. Physiol. 59,325 -347.[CrossRef][Medline]
Perry, S. F. and Goss, G. G. (1994). The effects of experimentally altered gill chloride cell surface area on acid-base regulation in rainbow trout during metabolic alkalosis. J. Comp. Physiol. B 164,327 -336.
Perry, S. F. and Laurent, P. (1993). Environmental effects on fish gill structure and function. In Fish Ecophysiology (ed. J. C. Rankin and F. B. Jensen), pp.231 -264. London: Chapman and Hall.
Rombough, P. J. (1999). The gill of fish larvae. Is it primarily a respiratory or an ionoregulatory structure? J. Fish Biol. 55,186 -204.[CrossRef]
Sakamoto, T., Yokota, S. and Ando, M. (2000). Rapid morphological oscillation of mitochondrion-rich cell in estuarine mudskipper following salinity changes. J. Exp. Zool. 286,666 -669.[CrossRef][Medline]
Steinhardt, R. A., Bi, G. and Alderton, J. M. (1994). Cell membrane resealing by a vesicular mechanism similar to neurotransmitter release. Science 263,390 -393.[Medline]
Tsai, J. C. and Hwang, P. P. (1998). The wheat germ agglutinin binding sites and development of the mitochondria-rich cells in gills of tilapia (Oreochromis mossambicus). Fish Physiol. Biochem. 19,95 -102.[CrossRef]
Van Der Heijden, A. J. H., Van Der Meij, J. C., Flik, G. and Wendelaar Bonga, S. E. (1999). Ultrastructure and distribution dynamics of chloride cells in tilapia larvae in fresh water and seawater. Cell Tissue Res. 297,119 -130.[CrossRef][Medline]
Van Der Heijden, A. J. H., Verbost, P. M., Eygensteyn, J., Li,
J., Wendelaar Bonga, S. E. and Flik, G. (1997).
Mitochondria-rich cells in gills of tilapia (Oreochromis mossambicus)
adapted to fresh water or seawater: quantification by confocal laser scanning
microscopy. J. Exp. Biol.
200, 55-64.
Wilson, J. M. and Laurent, P. (2002). Fish gill morphology: inside out. J. Exp. Zool. 293,192 -213.[CrossRef][Medline]
Witters, H., Berckmans, P. and Vangenechten, C. (1996). Immunolocalization of Na+, K+-ATPase in the gill epithelium of rainbow trout, Oncorhynchus mykiss. Cell Tissue Res. 283,461 -468.[CrossRef][Medline]