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Am J Physiol Regul Integr Comp Physiol 276: R363-R372, 1999;
0363-6119/99 $5.00
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Vol. 276, Issue 2, R363-R372, February 1999

Isolation of viable cell types from the gill epithelium of Japanese eel Anguilla japonica

Chris K. C. Wong and D. K. O. Chan

Department of Zoology, The University of Hong Kong, Hong Kong


    ABSTRACT
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

High-purity viable cells with low mitochondria (pavement cells) and mitochondria-rich content (chloride cells) were successfully isolated from the gill epithelium of Japanese eels, using three-step Percoll gradient low-speed centrifugation. Cytochemistry (silver staining for chloride, rhodamine-123, and Mitotracker for mitochondria and actin/spectrin immunofluorescence) and scanning electron microscope images were used to identify the cell types in the gill epithelium of the eel. Pavement cells were isolated at 97 and 98% purity for freshwater- and seawater-adapted eels, respectively, and chloride cells were obtained at 89 and 92% purity. The enzymatic activities of the isolated cells were determined. Na+-K+-ATPase, Mg2+-ATPase, and succinate dehydrogenase were found mainly in the chloride cell. Alkaline Ca2+-ATPase and low- and high-affinity Ca2+-ATPase were about twice as high in the chloride cell compared with the pavement cell. Transfer of eels to seawater resulted in enlargement of chloride cell sizes and significant increases in Na+-K+-ATPase, Mg2+-ATPase, and succinate dehydrogenase activities, while all Ca2+-ATPases declined by ~60-80%. This is the first report demonstrating the successful isolation of freshwater chloride cells and also an exclusive method of getting high-purity seawater chloride cells. The isolated cells are viable and suitable for further cytological and molecular studies to elucidate the mechanisms of ionic transport.

chloride cell; pavement cell; Percoll; mitochondria; adenosinetriphosphatase


    INTRODUCTION
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

THE FRESHWATER EEL actively takes up Cl-, Na+, and Ca2+ from the water to balance out losses of electrolytes in the urine (2). In seawater, the eel achieves electrolyte homeostasis by actively secreting Cl- and Na+ (21) into the water, but the role of the gill in Ca2+ homeostasis is not known. The seawater-adapted kidney excretes high levels of Ca2+ (3), which can account for the elimination of the Ca2+ absorbed along with drinking.

A series of experiments has been used to study ionic transport across the gill epithelia. Using the perfused head and opercular membrane preparations, freshwater chloride cells have been implicated as the possible site of Ca2+ uptake (23, 25, 29), while seawater chloride cells were responsible for Cl- extrusion (34). The cellular location of Na+ and Cl- uptake in freshwater environment is still controversial (27). Nevertheless, it is believed that chloride cells should play a principal role in it.

The "chloride-secreting cell" in the gill epithelium was first shown by Keys and Willmer (19) to undergo marked morphological changes following transfer of the eel from fresh to sea water. Since then, the chloride cell has been described in other teleost fishes and shown to respond to a rise in salinity with an increase in number and/or size (15, 31, 35) and to undergo marked ultrastructural changes, including increased mitochondria content, proliferative extension of the tubular system by invagination the basolateral plasma membrane (31, 35), and enlargement of the apical pit into which Cl- was excreted (6). Na+-K+-ATPase/ouabain binding sites of the gill tissue also increased (8, 17, 18, 33). Although a great deal of information has been gathered, the cellular mechanisms involved in ion transport in freshwater and seawater are still not fully understood. One inherent difficulty is the complexity of the gill epithelium, making it impossible to assign a particular function to a specific cell type. The gill is certainly involved in gaseous exchange, acid-base balance, and nitrogen excretion. Some of these actions no doubt take place across the "pavement" epithelial cells, but suggestions have been made that the chloride cell may also be involved.

A relatively enriched chloride cell fraction had been isolated from the gill epithelia of Anguilla japonica (16), A. anguilla (24, 33), Lagodon rhomboides (15), and Opsanus beta (28). However, only limited biochemical and physiological studies had been made on these cells. The present study was conducted to develop methods to isolate and characterize viable chloride cells of high purity from the gills of Japanese eels that had been adapted to different salinities for varying periods of time.


    MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

Animals

Japanese eels (Anguilla japonica), weighing between 500 and 600 g, were reared unfed in fiberglass tanks supplied with charcoal-filtered aerated tap water for at least 3 wk. The fish were then transferred to 50% seawater and full seawater and killed at predetermined intervals from 1 day to 3 wk.

Scanning Electron Microscopy

The excised gill arch (n = 12) was fixed overnight in 2.5% glutaraldehyde in Hanks' balanced salt solution (HBSS) at room temperature. After dehydration in ethanol and critical point drying with CO2, the tissue was mounted onto aluminum stubs and coated with gold palladium. The preparation was then examined under a Leica Steroscan-440 scanning electron microscope.

Cytochemical Characterization

Chloride. Gill arches (n = 5) were fixed briefly in buffered Formalin and then immersed in a solution of 1% AgNO3 and 2% HNO3 overnight. The tissues was rinsed in distilled water, blotted dry on filter paper, and refixed in Bouin's fluid. Five-micrometer wax sections were prepared and examined (6).

Mitochondria. Mitochondria in fresh gill tissues (n = 6) were stained by incubation with 1 µM rhodamine-123 (Calbiochem) or Mitotracker (CMTMRos-H2) (Molecular Probes) for ~45 min at room temperature. After washing in 10 mM PBS, the preparation was examined in a laser confocal microscope using an argon light source (Bio-Rad, MRC-600). Tissues stained with Mitotracker were postfixed in 4% (wt/vol) paraformaldehyde in 10 mM PBS (pH 7.2) at 4°C overnight, and the whole tissue was examined under the laser confocal microscope.

Actin and spectrin. Bouin-fixed wax sections of the gill tissue (n = 8) were dewaxed, rehydrated in graded ethanol, and rinsed in PBS. The staining procedure involved pretreatment of tissue sections with 10% normal goat serum in PBS to reduce nonspecific staining, followed by a 1-h incubation at room temperature with antiserum [rabbit anti-actin (1:100) or mouse anti-spectrin (1:500)] and a 1-h incubation with goat anti-rabbit or mouse IgG coupled to FITC (1:40) and then mounted in a 9:1 mixture of glycerol and PBS and examined by the confocal microscope. The slides were washed 3 × 15 min in the PBS after each antiserum application. Control procedures included the application of preimmune rabbit serum.

Acid mucoid. Acid mucoid material was stained with 1% toluidine blue to detect metachromasia.

Cell Isolation

The eels (n = 46) were anesthetized by tricaine methanosulfonate (1-2%). Gills were perfused with a buffered saline (in mM: 130 NaCl, 2.5 KCl, 5 NaHCO3, 2.5 glucose, 2 EDTA, 10 HEPES, pH 7.0), to remove blood cells. Gill arches were excised and washed. Epithelia were scrapped off from underlying cartilage with a glass slide into Ca2+, Mg2+-free HBSS (Sigma). The scrapings were dispersed by passing through two stainless steel filters with mesh size of 104 and 73.7 µm. They were then digested by 1.25 mg/ml collagenase (Sigma type I-A) and 2 mg/ml hyaluronidase (Sigma type I-S) at room temperature for 30-45 min. The cell suspension was washed by Ca2+, Mg2+-free HBSS and finally resuspended in 1.06 g/ml Percoll solution. Initially, a step gradient of 1.09, 1.08, 1.07, 1.06, 1.05, 1.04, and 1.03 g/ml was prepared and was centrifuged at 2,000 g, 15°C, for 15-20 min. For subsequent studies, a three-step gradient of 1.09, 1.06, and 1.03 g/ml was used instead. Isolated cell fractions were concentrated by centrifugation, and the pellet was processed for histochemical and scanning electron microscopic examination as in fixed tissues.

Cell Sizing and Counting

Isolated cells were counted and sized using a Coulter Multisizer II, with an orifice tube 70 µm in diameter and with isoton II as electrolyte. The cell count signal was the change of conductance of the electrolyte induced by particle resistance. Isoton II was used as blank, and calibration was carried out with monodiameter particles (PDVB latex 5.06 µm, Coulter). An aperture coincidence correction was below 2%.

Enzymatic Assays

Isolated cells were homogenated in a buffer (250 mM sucrose, 50 mM imidazole-HCl, 2 mM sodium EDTA, and 2 mM beta -mercaptoethanol, pH 7.2). The homogenate was centrifuged at 300 g for 10-15 min. The supernatant was further centrifuged at 14,000 g for 20 min. The final supernatant was stored at -76°C and assayed for Na+-K+-ATPase, Mg2+-ATPase, and Ca2+-ATPase. Protein content was measured by Bio-Rad protein assay kit.

Na+-K+-ATPase and Mg2+-ATPase. The activity was measured in a 2.5-ml medium containing 100 mM NaCl, 0.1 mM H2-EDTA, 12.5 mM KCl, 5 mM MgCl2, 0.4 mM NADH, 2 mM phosphoenolpyruvate, 1 mM ATP, 30 µl pyruvate kinase-lactate dehydrogenase (Sigma), and 30 mM imidazole, pH 7.4. The oxidation of NADH was monitored at 340 nm, which directly measured the rate of hydrolysis of ATP. The Na+-K+-ATPase activity was the difference in the enzymatic activity measured in the presence of 1 mM ouabain and a ouabain-free medium.

Alkaline Ca2+-ATPase. The cell extract was incubated in a medium containing 5 mM Ca2+, 70 mM NaCl, and 20 mM Tris-Cl buffer, pH 8.2, in a final volume of 0.6 ml. The mixture was prewarmed for 10 min at 25°C, and the reaction was started by addition of Na2-ATP (final concentration of 5 mM). After 1 h, 0.8 ml of a 10% TCA-2% ascorbic acid mixture was added to stop the reaction. The Ca2+-ATPase was the difference between the inorganic phosphate liberated in the presence of Ca2+ or EGTA. Inorganic phosphate was determined by the method of Fiske and Subbarow (11).

Low- and high-affinity Ca2+-ATPase. The standard assay incubation contained 200 µM EGTA, 10-2,400 µM CaCl2 (0.0035 µM-1.30 mM free Ca2+), 1 µM Na2-ATP, 20 mM sodium azide, and 12.5 mM Tris-1,4-piperazinediethanesulfonic acid, pH 7.4, at 25°C (30). After incubating for 1 h, the reaction was stopped as for alkaline Ca2+-ATPase. The enzyme activity was obtained by subtracting values obtained with chelator alone from those obtained with Ca2+ plus chelator.

Succinate dehydrogenase. Cells were homogenated and centrifuged at 300 g for 10-15 min. Supernatant was stored at -76°C for subsequent assay. The enzyme activity was measured in a 1-ml medium containing 50 mM K2HPO4-KH2PO4, 50 mM sodium succinate, 25 mM sucrose, 1 mg/ml 2-(p-iodophenyl)-3-p-nitrophenyl-5-phenyltetrazolium hydrochloride, pH 8.0. Incubation was carried out at room temperature until appropriate pink coloration developed. The reaction was stopped by addition of 100 µl 40% TCA. Reaction products were extracted by 4 ml ethyl acetate overnight at 4°C. The activity was determined in organic phase by measuring an absorbance at 490 nm (26).

Determination of Viability

A trypan blue exclusion test was employed to assess viability of the isolated cells. One drop of cells was incubated with one drop of 0.4% trypan blue for ~5 min. Stained cells and total cells were counted using a hemocytometer.

Statistical Analysis

All data are represented as means ± SE. Statistical significance was tested by Student's t-test and ANOVA depending on the number of means. The level of significance was set at P < 0.001.


    RESULTS
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

Gill Epithelial Morphology

The external surfaces of the primary and secondary gill lamellae were covered by pavement cells, which showed concentrically arranged ridges on their surfaces (Fig. 1, A and B). Mucous cells were quite rare and occurred on the leading (rostral) edge of the filament where the efferent vascular elements were located.


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  		Fig. 1.   Scanning electron micrographs showing distribution of chloride cells (arrowheads) on filamental and lamellar epithelia of freshwater (A)- and seawater (B)-adapted eels. Pavement cells and chloride cells are characterized by microridge (single arrows) and microvilli (double arrows) appearance, respectively. C illustrates that apical area (single arrow) was shared by 2 seawater chloride cells (*). Scale bars represent 3 µm.

Chloride cells were identifiable by their apical pit extrusions extending out between pavement cells, and were located mainly on the trailing (caudal) edge or afferent filamental surface, at the base of the secondary lamellae, and in the interlamellar space. In freshwater gills (Fig. 1A), chloride cells tend to occur singly, whereas in seawater gills (Fig. 1, B and C), two or three chloride cells shared a common depression between pavement cells. The crypt openings measured ~4.34 ± 0.92 µm × 1.98 ± 0.41 µm in freshwater eels. In seawater-adapted eels, the crypt openings were reduced and the longer diameter measured only 3.43 ± 0.6 µm (n = 41, P < 0.05).

In the cross section, the chloride cell could be easily identified by the dense precipitation of silver in the apical pit (Fig. 2A). Inside the cell, there was also a graded concentration of silver precipitates, with densities increasing toward the apical pit (n = 72, r = 0.847, P < 0.05). The silver-impregnated cell was also positive for anti-actin-FITC immunofluorescence. The FITC-green fluorescence was located in the whole cell (total pixel = 0.27 ± 0.03 × 106, n = 48) except the nuclear region (Fig. 2B). Chloride cells also showed intense mitochondria staining (Fig. 2C). Red blood cells showed intense staining for spectrin (Fig. 2D). The mucous cell stained with toluidine blue to yield a pink color (metachromasia), but did not stain with mitochondria stains, silver, actin, or spectrin. The pavement cell lining of the gill surface did not stain with silver, mitochondria stains, actin (total pixel = 0.07 ± 0.002 × 106, n = 38), or spectrin.


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  		Fig. 2.   A: deposition of silver precipitates in apical pits (arrowheads) of chloride cells, with densities increasing toward the mucosal side. B: immunocytochemical localization of actin filament in chloride cells (arrows) at the bases of lamellae and in the interlamellar region. C: mitochondria staining of chloride cells (arrows) localized in filamental and lamellar epithelia. D: immunocytochemical localization of spectrin in red blood cell (arrow). Scale bar in A represents 10 µm; scale bars in B-D represent 50 µm.

Step Percoll Gradient and Cell Identification

Seven-step Percoll gradient centrifugation yielded cells in eight layers. According to the distribution of cell size, there were four main populations of cells. The top layer (density < 1.03 g/ml) consisted of mucous (showing toluidine blue metachromasia) and dead cells (trypan blue positive). Mucous cells had large and numerous storage vesicles in the scanning electron microscope image (Fig. 3G). Layers 2-4 (1.03 g/ml < rho  < 1.06 g/ml) were composed of cells corresponding to a cell diameter (Coulter Multisizer) of 8-10 µm, which accounted for 87-100% of total cells in this fraction for freshwater eels and 66-98% for seawater-adapted eels. The remaining cells had diameters of 10-16 µm. There was only a very small percentage of contamination of mucous cells. The predominant cells recovered in layers 2-4 did not stain for actin, spectrin, or silver and had low mitochondria content (total pixel = 0.55 ± 0.04 × 106, n = 32). Scanning electron microscopy confirmed that these were pavement cells that lined the gill surface and showed characteristic ridges on their cell surface (Fig. 3, A and B). Layers 5-7 (1.06 g/ml < rho  < 1.09 g/ml) were composed of cells with diameter of 10-16 µm. In gills of freshwater eels, these cells had a mean size of 11.5 µm and accounted for 86-98% of harvested cells in these fractions (Fig. 3, C and D). In gills of seawater eels, the mean size was 15.5 µm and accounted for 85-98% of harvested cells. The predominant cell had a granular appearance and stained for mitochondria (freshwater cell: total pixel = 2.35 ± 0.18 × 106, n = 41; seawater cell: total pixel = 4.21 ± 0.13 × 106, n = 86, P < 0.001) (Fig. 3, E and F), silver, and actin. They were "chloride cells." The apical pit was well preserved in isolated cells. On the basis of the mitochondria density, two types of chloride cells could be discerned.


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  		Fig. 3.   Scanning electron micrographs showing ultrastructure of isolated pavement cells (A), freshwater chloride cell (C), seawater chloride cell (E), mucous cell (G), and red blood cells (H). Confocal microscope photomicrographs show mitochondria staining in both freshwater (D) and seawater (F) chloride cells, whereas pavement cells (B) show negative result. Single and double arrows in F represent 2 types of chloride cells with different mitochondria density. Scale bars in A, C, E, G, and H represent 3 µm; scale bars in B, D, and F represent 10 µm.

Layer 8 (density > 1.09 g/ml) was composed of cells that were ~5 µm in diameter and were identified as red blood cells by their spectrin staining and their ultrastructure (Fig. 3H).

For higher efficiency and yield, a three-step gradient (1.03, 1.06, and 1.09 g/ml) was designed. Again, the top cell layer consisted of mucous cells and dead cells. Layer 2, at the interface between 1.03 and 1.06 g/ml, consisted predominantly of pavement cells (8-10 µm diameter), with the cell purity measuring 97% for freshwater eels and 98% for seawater eels. Layer 3, at the interface between 1.06 and 1.09 g/ml, consisted mainly of chloride cells (10-16 µm), with cell purity of ~89% for freshwater eels and 92% for seawater eels.

Multisizer counting demonstrated that layer 3 cells increased in mean size from 11.5 to 15.5 µm within 4 days after seawater transfer (Fig. 4). Furthermore, for the same weight of starting materials, the yield of layer 3 cells from the seawater-adapted eel (2.95 ± 0.12 × 106) was ~10-fold higher than that of freshwater eel (0.31 ± 0.02 × 106). In contrast, no change in size was recorded for cells in layer 2 following transfer to seawater.


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  		Fig. 4.   Changes in sizes of layer 2 and layer 3 cells isolated from gill epithelia of eels following transfer from freshwater to seawater. * P < 0.05 compared with corresponding value in freshwater eel.

Enzymatic Assays

Enzymatic analyses were based on the cells separated by three-step Percoll gradient centrifugation. In both freshwater and seawater gills, layer 3 cells had significantly higher Na+-K+-ATPase activities than the corresponding layer 2 cells (6-fold difference in freshwater eel and 30-fold difference in seawater eel, P < 0.001 in both cases) (Table 1). Thus adaptation to seawater resulted in a 16-fold increase of Na+-K+-ATPase activity in the layer 2 cells (P < 0.001) and an 84-fold increase in the layer 3 cells (P < 0.001). For Mg2+-ATPase activity, there was no significant difference between freshwater layers 2 and 3 cells. Following adaptation to seawater, the enzyme activity increased significantly in layer 3 cells (40-fold increase).

                              
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  		Table 1.   Transport ATPase in gill cells isolated by a three-step Percoll gradient (layer 2 and layer 3): enzymatic activity

The enzyme alkaline Ca2+-ATPase was highest in freshwater layer 3 cells, at a level more than twice that in layer 2 cells. Following full adaptation to seawater, the enzyme activity was reduced to ~25% in both groups. For low- and high-affinity Ca2+-ATPase, the dependence of Ca2+-ATPase activity on Ca2+ concentration revealed a relationship representing the sum of two saturable components (Table 2). A high-affinity component was evident below 0.7 µM. Between 0.7 and 1.462 µM, there was minimal change in reaction rates. Above 5 µM, the ATPase activity again rose sharply. The affinity for Ca2+ was determined by presenting the results on an Eadie-Hofstee plot. In freshwater, the Km and Vmax of the low-affinity Ca2+-ATPase of low-mitochondria cells and mitochondria-rich cells were 13.14 µM Ca2+ and 11.93 nmol Pi · mg-1 · min-1 and 9.87 µM Ca2+ and 48.43 nmol Pi · mg-1 · min-1, respectively. For the high-affinity Ca2+-ATPase, the Km and Vmax of the low-mitochondria cells and mitochondria-rich cells were 0.043 µM Ca2+ and 2.57 nmol Pi · mg-1 · min-1 and 0.087 µM Ca2+ and 12.1 nmol Pi · mg-1 · min-1, respectively. Following adaptation to seawater, there were significant declines in the activities of both low- and high-affinity Ca2+-ATPase. The Km values were unaltered in most cases, except for the low-affinity enzyme in the layer 3 cells (chloride cells), where seawater adaptation increased the Km from 9.87 to 31.3 µM.

                              
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  		Table 2.   High- and low-affinity Ca2+-ATPase in the gill cells of the Japanese eel adapted to freshwater or seawater

Activities of succinate dehydrogenase (Table 3) were higher in layer 3 than the corresponding layer 2 cells (1.2-fold in freshwater and 3.8-fold in seawater, P < 0.001). The enzymatic activities increased in both layers following adaptation to seawater. Layer 3 cells had eightfold higher succinic dehydrogenase activity than those in freshwater.

                              
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  		Table 3.   Succinate dehydrogenase in gill cells isolated by a three-step Percoll gradient (layer 2 and layer 3): enzymatic activity and ratio of transport ATPases to succinic dehydrogenase activity

Trypan blue exclusion test showed that cell viability was ~90% for the isolated layer 2 cells and 92% for the layer 3 cells. Transfer to seawater had no effect on the proportion of viable cells harvested, although the total number of layer 3 cells harvested increased dramatically.


    DISCUSSION
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

Several isolation methods have been used to harvest chloride cells from the fish gill in the past 20 years, such as velocity sedimentation (24), step dextran gradient centrifugation (16), step Ficoll gradient centrifugation (15, 33), and continuous Percoll gradient centrifugation (28). Some methods were applied to seawater fish only and some were applicable to both freshwater and seawater fish. These results strengthen the view that the chloride cells in branchial epithelia are important in mediating the processes of ionic transport. However, the methods produced the isolated cells with either low purity (45-76%) and/or low cellular integrity (16, 33), which severely limit future cytological and physiological studies. In the present study, we make a great improvement on the methodology, which minimizes the cellular damage and maximizes the purity of the isolated cells. The purity of seawater chloride cells was 92%, which is better than that obtained by Naon and Mayer-Gostan (24) (76%), Hootman and Philpott (15) (60%), Sargent et al. (33) (45%), Kamiya (16) (enriched fraction), and Perry and Walsh (28) (enriched fraction). In addition, this is the first report demonstrating the successful isolation of freshwater chloride cell with purity over 89%. Furthermore, the isolated cells could be used in other functional studies (5), indicating a breakthrough in this area of study.

Among all the gradient forms, Percoll has been selected in this study because of its low viscosity and adjustable osmolarity, making it superior in terms of the maintenance of the cellular integrity. In combination with the step-gradient method, the present results showed that separation could be carried out under a relatively mild gravitational force. To get rid of blood cells, perfusing the gill with heparinized saline was essential. The scrapped epithelium was sieved to remove large clumps and filamental fragments. The high reaggregation tendency of the gill cells produced a problem in cell isolation. Trypsinization tends to produce a viscous complex of isolated cells. To minimize this effect, our preliminary trials showed distinct advantages to omitting trypsin as the tissue dissociation enzyme. A combination of collagenase and hyaluronidase was found to yield a suspension of free single cells. In early studies, a seven-gradient centrifugation was designed. Eight separated layers of cells were obtained. Comparing the freshwater and seawater sample, we noticed that there was an increase in the yield of large-size and high-density cells in the latter. However, the seven-step gradient had low cell recovery, and a longer centrifugation time was required to reach equilibrium. Some of the cells would span among the gradient solution rather than at the interface of the gradient. The three-step gradient centrifugation was found to be more satisfactory. The cells isolated in layer 2 and layer 3 reached in excess of 95% homogeneity, and a high percentage in excess of 90% were shown to be viable by the trypan blue exclusion test.

Colocalization of silver and mitochondria staining provided unequivocal identification of the mitochondria-rich cell as the chloride cell. The high content of actin fibers in the chloride cell probably played an essential role in maintaining cell shape and apical pit architecture for this ion-transporting cell. All three staining procedures confirmed that the chloride cell could be isolated in high purity in the layer 3 fraction of the three-gradient Percoll gradient centrifugation. Biochemical data on Na+-K+-ATPase and succinic dehydrogenase also provided support for the identity of this cell. Among the layer 3 cells, some had a very dense concentration of mitochondria whereas others had distinctly fewer mitochondria. Thus two types of chloride cells coexisted in both freshwater- and seawater-adapted eels (unpublished results). These could correspond with the two-cell type previously described in many teleost species on the basis of transmission electron microscopy studies (32, 35). Because pavement cells were not positively stained with any test in this study, surface ridges provided the only reliable criterion for the identification of these cells after isolation. The presence of concentric ridges on the surface of the pavement cell implied an essential function such as increasing surface area for respiration.

Apart from mitochondria-specific fluorescence dyes (rhodamine-123 and Mitotracker), Mg2+-ATPase (7) and succinic dehydrogenase (33) were also located exclusively in the mitochondria and hence provide a marker for the mitochondria-rich chloride cell. In freshwater eels, chloride cells had 16 and 22% more succinic dehydrogenase activity and Mg2+-ATPase activity per milligram cellular protein, respectively, compared with the pavement cells. Following adaptation to seawater, both enzymes increased about twofold in the pavement cells. However, in the chloride cells, succinic dehydrogenase activity increased 8-fold while Mg2+-ATPase activity increased 40-fold. Thus the seawater chloride cell appeared to be more efficient in generating ATP than succinic acid oxidation. This shift suggested that seawater adaptation actually involved not only an increase in the cell size for the chloride cell but also significant reorganization of the structural units making up the mitochondria inner membranes, boosting its efficiency in ATP production.

The Na+-K+-ATPase enzyme has been shown to be associated exclusively with the basolateral plasma membrane and its invaginated system of tubules. The enzyme increased by 84-fold in the chloride cell following adaptation to seawater; this must have arisen from the proliferative extension of the intracellular tubule system, which must form an essential component in the Na+-extrusion mechanism that operates in the seawater-adapted chloride cell. The results of multisizer counting clearly demonstrated that entry to seawater caused significant increases in the size and number of chloride cells that took place over several days. The increases in size and the yield of chloride cells during seawater transfer provide unequivocal proof of the importance of this cell type in ionic transport in seawater. Cortisol has been shown to be essential in triggering seawater adaptation and Na+ extrusion in the teleost fish (22). Indeed, cortisol injection into eels maintained in freshwater stimulated production of the seawater-type chloride cell (high mitochondria autofluorescence, large cell size) within 4-6 days (4). On a proportional basis, using the freshwater system as the baseline, the increase in Na+-K+-ATPase activity following seawater adaptation was much more significant compared with the increase in mitochondria enzymes, e.g., succinic dehydrogenase and Mg2+-ATPase.

Chloride cells had higher levels of all three Ca2+-ATPase enzymes compared with the pavement cells. All three enzymes decreased to about the same extent in both chloride cells and pavement cells following transfer of the eel from freshwater to seawater. Considering that there was ~10-fold increase in the yield of chloride cells after adaptation to seawater, the total Ca2+-ATPase in the gill epithelium could actually increase. Ma et al. (20) first reported there was no distinct difference in gill total alkaline Ca2+-ATPase activity when freshwater rainbow trout were adapted to seawater. Burdick et al. (1), on the other hand, demonstrated a higher gill alkaline Ca2+-ATPase activity in seawater-adapted killifish. Ho and Chan (14), working on the Japanese eel, reported a significant rise in serum Ca2+ concentration during the initial 2-4 days following transfer to seawater, but this declined and remained regulated from day 6 onward when branchial alkaline Ca2+-ATPase also increased and remained high as the eel fully acclimated to the seawater. However, Fenwick (10) reported that alkaline Ca2+-ATPase activity was higher in gills of freshwater-acclimated American eel than in the gills of seawater-acclimated eel. These contradictory results cast serious doubts on the direct role alkaline Ca2+-ATPase plays in the active transepithelial transport of calcium through the gills. A better understanding on the role of this enzyme must depend on direct measurements of Ca2+ transport itself.

Fenwick (9) and So and Fenwick (36) measured the rate of influx of 45Ca2+ and the activity of alkaline Ca2+-ATPase in perfused isolated American eel gills and found a direct positive correlation. Additionally, stanniocalcin, which decreased uptake of Ca2+ from the water, also suppressed branchial alkaline Ca2+-ATPase activity (20). The pH optimum for this Ca2+-ATPase was ~8.0 (9, 14, 20), and the Km value was in the 0.4-0.6 mM range and was mersalyl sensitive (14). However, the subcellular location of this low-affinity and high-capacity Ca2+-ATPase was microsomal (14). In the present study, changes in the alkaline Ca2+-ATPase levels in both chloride and pavement cells were completely dissociated from those of mitochondria enzymes, thus supporting the microsome origin of this enzyme. It had been pointed out by later workers that to function as a Ca2+ extrusion pump to transport Ca2+ out of the cell, the Ca2+-ATPase must be capable of being activated by concentrations of ionized Ca2+ in the micromolar range. Therefore, the low-affinity Ca2+-ATPase may not be involved in the maintenance of submicromolar intracellular Ca2+ concentrations found in resting cells. Flik et al. (12) reported the presence of a high-affinity Ca2+-ATPase activity in the eel gill plasma membranes and found a Km value for Ca2+ of 0.22 µM. In addition, induction of this high-affinity Ca2+-ATPase activity in eel gill plasma membranes was observed after treatment of eels with prolactin, which suggested an involvement of this enzyme in the active uptake of Ca2+ from the freshwater (13).

In the present study, both alkaline and neutral low- and high-affinity Ca2+-ATPase activities were high in the freshwater chloride cell and were reduced by the same extent following adaptation to seawater. This synchronized pattern of changes suggested that these enzymes might serve as a package, with the low-affinity enzyme serving for high-capacity transport associated with the accumulation of intracellular ionized Ca2+ into a microsomal Ca2+ "sink." Final reduction of Ca2+ concentration would be achieved by the high-affinity, low-capacity Ca2+-transport ATPase. Recently, we have shown that after labeling isolated eel chloride cells with the calcium-fluorescence dye fluo-3 or the ratiometric dye Indo-1, the highest intensity of Ca2+ fluorescence was initially polarized to a small spot ~<FR><NU>1</NU><DE>6</DE></FR> the diameter of the cell and then spread to the rest of the cell (5), suggesting the entry of Ca2+ through the apical pit of the cell.

In conclusion, we demonstrate clearly the method of pavement and chloride cells isolation and also illustrate the cytological and biochemical changes of the cells associated with external salinities. The consequence of increased Ca2+-ATPase in freshwater chloride cell or increased Na+-K+-ATPase in seawater chloride cell suggested an importance of the cell in mediating Ca2+ uptake and Na+ extrusion, respectively.

Perspectives

In the past 60 years, our understanding on the gill ion transport has relied on the electrophysiological, morphometric, and biochemical studies of the whole gill preparations. These studies provide unequivocal evidence that strengthens the view of gill transepithelial transport associated with fish osmoregulation. However, the knowledge of the mechanisms of cellular ion transport (especially Na+, Cl-, and Ca2+ in freshwater and Na+ in seawater) are still limited. The studies described here on gill cell isolation represent only the first in a series of steps needed to further understand the pavement cells and chloride cells in ionic transport. The next step is to characterize the properties of calcium channels on the apical membrane of chloride cells by using patch-clamp technique. In addition, the decrease of intracellular Ca2+ level of freshwater chloride cell on the addition of stanniocalcin (5) suggested that stanniocalcin receptor was expressed in the cells. RNA fingerprinting could be applied to the isolated freshwater and seawater chloride cells to clone the receptor and also to distinguish the differential expression of particular functional genes. The technique developed in this study opens up a new approach to tackle the problems that have yet to be solved.


    ACKNOWLEDGEMENTS

This work was supported by the Research Grants Council, Hong Kong


    FOOTNOTES

Address for reprint requests: C. K. C. Wong, Dept. of Biology, Hong Kong Baptist Univ., Kowloon Tong, Hong Kong.

Received 31 December 1997; accepted in final form 30 September 1998.


    REFERENCES
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

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Am J Physiol Regul Integr Compar Physiol 276(2):R363-R372
0002-9513/99 $5.00 Copyright © 1999 the American Physiological Society


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