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MEMBRANE TRANSPORTERS, ION CHANNELS, AND PUMPS
Laboratory of Physiology, K.U. Leuven, Campus Gasthuisberg, Leuven, Belgium
Submitted 27 October 2005 ; accepted in final form 27 November 2005
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ABSTRACT |
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 TOP ABSTRACT METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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caveolae; osmolyte; epithelial cell; chloride channel
The ion channel that is responsible for the swelling-induced release of Cl– is known as the volume-regulated anion channel (VRAC) (also known as volume-sensitive organic osmolyte-anion channel or volume-sensitive outwardly rectifying channel). VRAC is a ubiquitously expressed plasma membrane anion-selective channel that has been extensively characterized at the biophysical and pharmacological levels (for reviews, see Refs. 15 and 23). VRAC seems to be a multipurpose ion channel (10, 24, 25), because in addition to its role in cell volume regulation the channel is also involved in apoptosis (36), setting of the membrane potential (47), cell proliferation (48), angiogenesis (19), and mechanosensation (22). Despite >10 yr of extensive work on VRAC, the molecular identity of VRAC remains enigmatic. Several candidates (e.g., P-glycoprotein, pICln, ClC-3, ClC-2) have been proposed, but there is no conclusive evidence or unanimous agreement for any of them at the moment (15, 23–25). A putative new Cl– channel family (hTTYH1–3) was recently identified, of which one member, hTTYH1, elicited a swelling-activated Cl– current after transient transfection in Chinese hamster ovary cells (41); however, the biophysical properties of hTTYH1 and VRAC are at first sight different. The ambiguity about the molecular identity of VRAC has hampered progress in the functional and structural characterization of VRAC. For example, it is not clear how VRAC relates to other swelling-activated transport pathways, e.g., those responsible for the swelling-induced release of organic osmolytes and ATP. Because the swelling-induced Cl– current through VRAC (ICl,swell) and the swelling-induced release of organic osmolytes share several characteristics in terms of pharmacology, kinetics, and activation parameters, it has been proposed that VRAC also functions as a pathway for organic osmolytes such as taurine, D-myo-inositol, and sorbitol (13). The estimated pore diameter of VRAC (
1.1 nm) is indeed compatible with permeability for taurine and polyols (9, 26). For this reason, VRAC is sometimes referred to as the volume-sensitive organic osmolyte anion channel. However, other groups (40, 42) have provided evidence that, at least in some cell types, VRAC and the taurine release pathway can be differentiated based on their pharmacology, kinetics, and sensitivity toward PKC. Similarly, it has been argued that swollen cells release ATP via VRAC (12), but alternative efflux pathways for ATP have been proposed: a maxi-channel that is different from VRAC (11, 30), Ca2+-dependent exocytosis (1) and ATP efflux via an ABC transporter [cystic fibrosis transmembrane conductance regulator (CFTR) or P-gp MDR1] (for review, see Ref. 35). Finally, one should keep in mind that these hypotheses (common vs. separate pathways) are not necessarily mutually exclusive. It is very well possible that VRAC is permeable to both taurine and ATP and that there exist additional taurine- or ATP-selective pathways, which may vary in expression levels from cell to cell.
Caveolae and lipid rafts are cholesterol- and sphingolipid-rich microdomains in the plasma membrane, which are implicated in a variety of cellular processes: signal transduction, clathrin-independent endocytosis/transcytosis, cellular cholesterol homeostasis and cell survival/apoptosis (for reviews, see Refs. 6, 29, 46). Caveolae form a subset of lipid rafts characterized by their morphology (caveolae form flask-shaped invaginations) and protein content (caveolae contain caveolin proteins) (37). Caveolin proteins not only function as scaffold proteins that induce and/or stabilize the caveolar invagination, but also are actively involved in regulating the activity of signaling proteins (G proteins, nonreceptor tyrosine kinases, and endothelial nitric oxide synthase) that are sequestered in caveolae (6). We have previously shown that caveolae/lipid rafts play a crucial role in VRAC expression and/or activation. First, hypotonic stimulation of caveolin-1-deficient cells, such as Caco-2, MCF-7, or T47D, triggers either no or a poor ICl,swell response, but current activation is restored by transient expression of caveolin-1 in these cells (45). Second, in calf pulmonary vascular endothelium cells, a vascular endothelial cell line with high endogenous caveolin-1 expression, expression of a dominant-negative caveolin-1 mutant represses VRAC activation (44). Third, transient expression of c-Src, a nonreceptor tyrosine kinase, downregulates VRAC activity in calf pulmonary vascular endothelium cells, provided that c-Src is targeted to lipid rafts/caveolae via a NH2-terminal dual acylation signal (43). Although these observations clearly establish a functional link between lipid rafts/caveolae and VRAC, they leave open a number of important questions. First, what is the underlying mechanism connecting lipid rafts/caveolae and VRAC? Second, is the effect of lipid rafts/caveolae restricted to VRAC or does it extend to other cell swelling-dependent processes such as the release of organic osmolytes?
In this study, we have addressed the role of caveolin-1, the principal scaffold protein of caveolae in nonmuscle cells, on swelling-induced release of taurine and ATP in a polarized intestinal epithelial cell line (Caco-2). The main outcome is that caveolin-1 exerts a polarized effect on the hypotonicity-induced release of taurine and ATP by selectively stimulating the basolateral release.
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METHODS |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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The human colon cancer cells Caco-2 (ECACC86010202) used in this study were either wild-type (WT) or stably transfected with the pcDNA3 expression vector containing the canine caveolin-1 cDNA. The cells were grown in DMEM containing 20% fetal bovine serum (Sigma, St. Louis, MO), 3.2 mM L-glutamine, 80 U/ml penicillin, 80 µg/ml streptomycin, and 0.8% MEM with nonessential amino acids. The growth medium for the stably transfected cells was supplemented with 800 µg/ml geneticin (G-418 sulfate; Invitrogen, Carlsbad, CA) to generate a polyclonal pool (cav1-K) from which the monoclonal cell lines (e.g., k14 and k37) were isolated by serial dilution cloning. For electrophysiological characterization, Caco-2 cells were directly seeded onto gelatine-coated coverslips at 5 x 103 cells per coverslip. The taurine and ATP release experiments were performed on a polarized monolayer obtained by seeding the cells onto permeable Anopore filters (pore size, 0.2 µm; Nunc Intermed, Roskilde, Denmark) at a density of 6 x 104 cm–2. The cells were kept in a humidified incubator (37°C and 9% CO2), and the growth medium was renewed twice a week. The polarized monolayers were used between days 7 and 10 for both taurine efflux experiments and ATP release experiments.
Immunoblot Analysis of Caveolin-1 Expression
Caco-2 cells (107) were lysed in a hypotonic buffer (1 ml) containing 25 mM Tris·HCl, pH 7.5, 20 mM NaCl, 2.5 mM EGTA, 0.5% (vol/vol) Nonidet P-40, and a protease inhibitor cocktail (1 mM phenylmethylsulfonyl fluoride and 0.1 µg/ml leupeptin). The lysate was centrifuged and the supernatant was stored at –20°C. Protein (50 µg) was separated on SDS-PAGE (12.5% acrylamide) and transferred onto a polyvinylidene difluoride microporous membrane (Immobilon, Millipore) by semidry electroblotting. Ponceau S staining was used to check the blotting efficiency. Caveolin-1 was detected with a commercially available monoclonal antibody (C37120 [GenBank] , Transduction Laboratories) diluted 1:1,000. Secondary antibodies were alkaline-phosphatase-conjugated goat anti-mouse IgG. Immunoreactive bands were visualized with the Vistra enhanced chemifluorescence detection kit (Amersham Biosciences/GE Healthcare) on a Storm 840 imager (Molecular Dynamics).
To study the subcellular distribution of caveolin-1, Caco-2 cells (5 x 106) were harvested by scraping in ice-cold 500-µl lysis buffer composed of (in mM) 25 Tris·HCl, pH 7.2, 50 NaCl, 90 mannitol, and 1 EGTA, supplemented with 1% Triton X-100, 2 mM DTT, 1 mM PMSF, 10 µg/ml leupeptine, and 10 µg/ml aprotinine. The lysate was adjusted to 45% sucrose and subjected to a sucrose density gradient centrifugation in a Beckman SW50.1 rotor at 250,000 g for 20 h at 4°C. The sucrose gradient was formed by layering 3 ml of 30% sucrose in lysis buffer and 1 ml of 5% sucrose in lysis buffer on top of the 1 ml lysate/sucrose 45%. After centrifugation, 500-µl fractions were collected from top to bottom (first 250 µl were discarded) and proteins were precipitated with 10% ice-cold trichloroacetic acid. Pellets were washed with acetone resuspended in Laemmli loading buffer, separated by SDS-PAGE (12.5% polyacrylamide), and transferred by semidry electroblotting onto polyvinylidene difluoride membrane (Immobilone, Millipore). Blots were stained with anti-caveolin-1 monoclonal antibody (C37120 [GenBank] ; Transduction Laboratories) as described above.
Electrophysiology
The standard extracellular solution was a modified Krebs solution containing (in mM) 150 NaCl, 6 KCl, 1 MgCl2, 1.5 CaCl2, 10 glucose, 10 HEPES, pH 7.4 and with NaOH. The osmolality of this solution as measured with a vapor pressure osmometer (Wescor 5500, Schlag, Gladbach, Germany) was 320 ± 5 mosmol/kgH2O. At the beginning of the patch-clamp recordings, the Krebs solution was replaced by an isotonic (ISO) Cs+ solution containing (in mM) 105 NaCl, 6 CsCl, 1 MgCl2, 1.5 CaCl2, 10 glucose, 10 HEPES, and 90 mannitol, pH 7.4 with NaOH (320 ± 5 mosmol/kgH2O). The 25% hypotonic solution was obtained by omitting 90 mM mannitol from this solution. The standard pipette solution contained (in mM) 40 CsCl, 100 Cs+-aspartate, 1 MgCl2, 1.93 CaCl2, 5 EGTA, 4 Na2ATP, and 10 HEPES, pH 7.2 with CsOH (290 mosmol/kgH2O). The concentration of free Ca2+ in this solution was buffered at 100 nM.
Currents were monitored with an EPC-7 patch-clamp amplifier (List Electronic, Lambrecht/Pfalz, Germany). Patch electrodes had DC resistances between 2 and 6 M
. Potentials were measured with reference to an Ag-AgCl electrode. Whole cell membrane currents were recorded using ruptured patches. Currents were sampled at 1-ms interval and filtered at 1,000 Hz. Capacitative and leak currents were compensated. Between 50% and 70% of the series resistance was compensated electronically to minimize voltage errors. The following voltage protocol was applied every 10 s from a holding potential of –25 mV: a step to –80 mV for 0.2 s, followed by a step to –100 mV for 0.1 s, and a 1.5-s linear voltage ramp to +100 mV. Experiments were performed at room temperature.
Electrophysiological data were acquired with pCLAMP 5.5 (Axon Instruments) and analyzed with WinASCD (provided by G. Droogmans; ftp://ftp.cc.kuleuven.ac.be/pub/droogmans/winascd.zip) and Origin 7.0 software (OriginLab, Northampton, MA). Time courses of the whole cell current were obtained by plotting the current at +80 or –80 mV during successive voltage ramps as a function of potential. Current-voltage relationships were obtained from the currents measured during the linear voltage ramp. ICl,swell was calculated by subtracting the basal current under ISO conditions from the maximal current during hypotonic stimulation.
Flux Experiments
Solutions. ISO solutions (330 mosmol/kgH2O) contained (in mM) 165 Na+, 6 K+, 1.5 Ca2+, 1 Mg2+, 10 glucose, 10 HEPES, and 166 Cl– (pH 7.4). Hypotonic solutions (170 mosmol/kgH2O) were prepared by the removal of 90 mM NaCl and hypertonic solutions (490 mosmol/kgH2O) by the addition of 87.5 mM NaCl. The osmolality of the solutions was verified with a cryoscopic osmometer (Osmomat 030, Gonotec, Berlin, Germany). 5-Nitro-2-(3-phenylpropylamino)benzoate (NPPB) and tamoxifen were purchased from Sigma (St. Louis, MO). ATP release was measured by using a luciferine-luciferase kit purchased from Sigma (FL-AAM). Fifty microliters per milliliter of this luciferin-luciferase assay (LL) were added to the solutions used to probe ATP release. This resulted in a final concentration of the following components: 0.5 mM MgSO4, 0.05 mM EDTA, 0.005 mM dithiothreitol, 2.5 mM tricine, 0.03 mM luciferin, 3.3 mg/l luciferase, and 50 mg/l bovine serum albumin. [1,2-3H]taurine (5–30 Ci/mmol) was purchased from Amersham Biosciences (GE Healthcare).
Taurine release. To measure taurine release, the cells were loaded with [3H]taurine. Both the basolateral and apical surfaces of the epithelium were incubated for 2 h in an ISO solution containing 0.3 µCi/ml of [1,2-3H]taurine. After being loaded, the tissues were rinsed with ISO solution to remove the remaining extracellular taurine. The monolayer was then removed from the filter cup and mounted in a two-compartment Ussing-type chamber with an exposed membrane area of 1.54 cm2. During the whole experimental protocol, both compartments were continuously perfused. The washout of the tracer from the Anopore filter was accelerated by stirring of the basolateral perfusate with a motor-driven magnetic stirring bar. The apical and basolateral perfusates were collected in scintillation vials that were replaced at 4-min intervals. [3H]taurine was measured by liquid scintillation counting. At the end of each experiment, the cells were lysed with a 5% (wt/vol) trichloroacetic acid solution and four additional samples were collected to determine the total amount of counts accumulated in the cells, which was, on average 1 x 105 counts per min. The data are presented as fractional taurine release and expressed as radioactivity released per minute in each collected sample as the percentage of the total amount in the cells at that time.
ATP release. The real-time recording of the ATP release, based on luciferin-luciferase (LL) bioluminescence, was enabled by the use of custom-built instrumentation and software. The setup used for basolateral ATP release was described by Jans et al. (14). ATP release from damaged cells was minimized by leaving the Anopore filter attached to the holder. The preparation was mounted in a light-tight container that separates the apical from the basolateral compartment. The ATP released in the basolateral perfusate was probed with the luciferin-luciferase (LL) mixture (50 µl/ml). Emitted photons were detected by a photon-counting tube (model H3460-04, Hamamatsu Photonics) that was positioned 2 cm from the apical surface of the epithelium. The light impulses were discriminated, prescaled, and counted with a personal computer-based 32-bit counter/timer board (model PCI-6601; National Instruments, Austin, TX). The number of impulses counted in 1-s time interval was monitored and graphically displayed. Because of the breakdown and the consumption of ATP and the LL reagent mixture, a pulse protocol was designed: the perfusion was interrupted during 90 s, which allowed the ATP to accumulate in the presence of the LL assay. During this short time period, breakdown and consumption of ATP and LL were negligible (14). The rate of ATP release was calculated by linear regression of the rising amounts of ATP during the stop-flow period of the pulse protocol. The apical ATP release was measured with a similar but adapted setup. Calibration of the ATP measurements was performed using the same setup with blank Anopore filters to mimic as much as possible the experimental conditions. The data are converted to the amount of ATP released by 1 µl of cell volume.
Protocols. The preparations were perfused at the apical and basolateral sides as follows: ISO solution (330 mosmol/kgH2O) for 24 min, hypotonic solution for 40 min (170 mosmol/kgH2O), ISO solution for 24 min. NPPB (100 µM) and tamoxifen (10 µM) were added to both sides of the epithelium. All experiments were performed at room temperature (22°C) in air-conditioned rooms.
Analysis
Pooled data are expressed as means ± SE from n cells. The significance between two data sets was tested using the Student's unpaired t-test. ANOVA was used to determine statistical differences of three or more data sets. Differences were considered significant at the level of P < 0.05.
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RESULTS |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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To study the effect of caveolin-1 on hypotonicity-induced transport processes, we used Caco-2 cells, either WT that express minimal amounts of caveolin-1 (45, 49), or caveolin-1-expressing Caco-2 cells obtained by stable transfection and clonal dilution. Caco-2 cells were stably transfected with a selectable caveolin-1 expression vector to generate cell lines permanently expressing caveolin-1. A polyclonal cell pool (cav1-K) was first generated from which monoclonal cell lines (k14 and k37) were isolated by clonal dilution. The expression of caveolin-1 was verified using Western blot analysis for the polyclonal pool (data not shown) and for the monoclonal cell lines (Fig. 1). Western blot analysis of total cell lysates showed a clear caveolin-1 signal in the k14 and k37 monoclonal cell lines, but no signal in WT Caco-2 cells (Fig. 1A). Caveolin-2 expression levels were comparable between WT and transfected Caco-2 cell lines. Caveolin-1 is a major structural protein of caveolae, which due to their lipid content (high cholesterol and sphingolipid content) can be biochemically isolated as detergent-resistant membrane fractions after Triton X-100 solubilization at 4°C and upward flotation in a sucrose density gradient centrifugation (37). As shown in Fig. 1B, Western blot analysis of detergent-resistant membrane fractions of k14 and k37 Caco-2 cells revealed strong caveolin-1 signals in the upper fractions of the sucrose gradient, which is consistent with the incorporation of the stably expressed caveolin-1 in detergent-resistant membrane fractions. Interestingly, a very faint caveolin-1 signal could be detected in detergent-resistant membrane fractions of WT Caco-2 cells indicating a (very) low level of endogenous caveolin-1 expression in WT Caco-2.
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-cyclodextrin or filipin did not affect anion secretion in colonic epithelium as measured by short-circuit current, again indicating that raft/caveola integrity is not required for proper CFTR functioning (17).
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Having verified the caveolin-1 expression in k14 and k37 Caco-2, we asked the question whether the functional effect of caveolin-1 was restricted to VRAC or whether other hypotonicity-induced transport processes, such as organic osmolyte release and ATP release, were also affected. Moreover, because Caco-2 cells grown as a confluent monolayer form a polarized cell layer, we investigated whether the effects of caveolin-1 were confined to either the apical or basolateral membrane. Because we could not discern any electrophysiological difference between k14 and k37 clone, these experiments were performed on the k14 cell line.
In a first series of experiments, we measured the efflux of taurine, an organic osmolyte that is released during hypotonic cell swelling (42). WT and k14 Caco-2 cells were grown on semipermeable filters for 7–10 days, after which they formed a confluent monolayer with a transepithelial resistance of 250 /cm2. Flux experiments were performed in a modified Ussing-type chamber, which allowed us to differentially sample the apical and basolateral taurine release before and during hypotonic cell swelling (application of a 170 mosmol/kgH2O extracellular solution at both the apical and basolateral side). As shown in Fig. 4A, the basolateral taurine efflux during hypotonicity was markedly higher in k14 than in WT Caco-2 cells, reaching a peak value of 17.1 ± 2.0 min–1 (achieved after 20 min of HTS) vs. 11.7 ± 1.3 min–1 (achieved after 24 min of HTS), respectively (n = 6; P < 0.05). Application of 100 µM NPPB, a well-characterized blocker of ICl,swell and taurine efflux (13, 32) inhibited the basolateral hypotonicity-induced taurine efflux both in WT and in k14 Caco-2 cells (Fig. 4A). Maximal basolateral taurine release in the presence of NPPB (100 µM) was achieved at a later time point during hypotonic stimulation, and the amplitude was reduced to 3.6 ± 0.6 min–1 for WT cells (n = 5; P < 0.05 vs. control WT) and to 6.1 ± 1.2 min–1 for k14 Caco-2 (n = 5; P < 0.05 vs. control k14). In contrast, 10 µM tamoxifen did not exert a statistically significant effect on the basolateral taurine release in WT or in k14 Caco-2 cells. An opposite effect of caveolin-1 expression was observed for the apical hypotonicity-induced taurine release (Fig. 4B). Hypotonic exposure induced a progressive increase in taurine efflux from the apical membrane in WT Caco-2, whereas in k14 Caco-2, the apical taurine efflux was reduced at all time points. Maximal apical taurine fluxes were, respectively, 7.7 ± 2.3 min–1 in WT Caco-2 vs. 5.3 ± 1.6 min–1 in k14 Caco-2 (n = 6 for both cell types), but these differences were not statistically significant at the level of 0.05. The apical HTS-induced taurine release was sensitive to NPPB, which nearly completely blocked the apical release, both in WT and in k14 Caco-2 cells (Fig. 4B). Whereas 10 µM tamoxifen hardly affected the basolateral taurine efflux, it nearly completely abolished the apical taurine efflux in both WT and k14 Caco-2 cells (Fig. 4D). Thus tamoxifen preferentially blocks the apical HTS-induced taurine efflux.
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In a final set of experiments, we investigated the apical and basolateral release of ATP during hypotonic stimulation. Again, the expression of caveolin-1 had opposite effects on the basolateral and apical hypotonicity-induced ATP release (see Fig. 6). Peak values for the basolateral ATP release at 10 min of hypotonicity were 0.32 ± 0.04 pmol/min for WT (n = 5) and 1.08 ± 0.22 pmol/min for k14 cells (n = 5; P < 0.05). In contrast, caveolin-1 expression did not stimulate the apical ATP release. Peak values for the apical release at 3-min hypotonicity were 0.49 ± 0.17 pmol/min for WT (n = 8) and 0.25 ± 0.08 pmol/min for k14 cells (n = 9; P > 0.05). Intriguingly, return to isotonicity did not trigger a transient increase in ATP release, as was observed for taurine.
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DISCUSSION |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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There are several plausible mechanisms by which to explain the effect of caveolin-1 on swelling-induced membrane transport as can be deduced from observations of other ion channels. First, caveolin-1 may be required for the proper plasma membrane location of the transporter or channel. This has been shown for TRPC1, a nonselective cation channel that might be involved in the store-operated Ca2+ entry in some cell types. Deletion of a NH2-terminal caveolin-1 binding site in TRPC1 prevented the proper location of the channel in the plasma membrane (4). Conversely, specific caveolin-1 deletion mutants led to the intracellular retention of TRPC1. Thus these data indicate that caveolin-1 plays a crucial role either in the delivery of TRPC1 to the plasma membrane or in its retention in the plasma membrane. Second, functional caveolae may be required for the proper assembly of signal transduction complexes. For example, both the ATP-sensitive K+ (KATP) channel and adenylate cyclase reside in caveolae in vascular smooth muscle, where they are assembled in a functional signal transduction complex that allows for efficient activation of the KATP channel by protein kinase A. Disruption of caveolae by cholesterol extraction with cyclodextrin reduced activation of the KATP current by protein kinase A (31). Similarly, it has been proposed that caveolae are required for the incorporation of TRPC1 in a signal transduction complex that also contains G
-subunits and inositol 1,4,5-trisphosphate receptors (18). Third, caveolin-1 can directly modulate the functional properties of an ion channel or transporter. For example, large-conductance, Ca2+-activated K+ channels in vascular endothelial cells or myometrial smooth muscle reside in caveolae, where they are kept in a repressed state by interaction with caveolin-1 (3, 50). Disruption of this interaction either by cholesterol extraction with cyclodextrin or by knocking down caveolin-1 via small interfering RNA deinhibits the ion channel. A fourth possibility is that caveolin-1 is part of the channel/transporter itself. Although this cannot be excluded a priori, there is at present no experimental evidence that caveolin-1 functions as a structural subunit of a membrane transporter or ion channel. Thus caveolin-1 and caveolae can exert either positive or negative effects on ion channels and membrane transporters. Stimulation is ascribed to a role for caveolin/caveolae either in plasma membrane delivery or in the formation of efficient signal transduction complexes. In contrast, inhibition results from a direct interaction between caveolin-1 and the partner protein, which as a result is clamped in an inactive or less active configuration. Insofar as this general scheme can be extrapolated to VRAC and the efflux pathways for taurine and ATP, it can be hypothesized that caveolin-1 expression either promotes the plasma membrane delivery of these transporters or mediates the organization of a swelling-activated signal transduction complex. Unfortunately, the lack of molecular data with respect to the identity of the swelling-induced transporters prevents us from differentiating these two scenarios.
A second conclusion from our data is that the effect of caveolin-1 on the swelling-induced release of taurine and ATP is polarized. Expression of caveolin-1 selectively stimulates the basolateral release but has no significant effect on the apical release. That caveolin-1 exerts an effect on basolateral processes is not surprising in view of its subcellular distribution in polarized epithelial cells. Immunofluorescence studies on kidney sections have shown that caveolin-1 is localized to the basolateral membrane of specific segments along the nephron (distal tubule cells, connecting tubule cells, and collecting duct principal cells) which coincides with the basolateral detection of caveolae using electron microscopy (5). Studies (34, 49) on polarized Madin-Darby canine kidney have shown that the apical membrane contains caveolin-1, but no caveolae, whereas the basolateral membrane contains both caveolin-1 and caveolae. Apparently, formation of caveolae requires the formation of heteromeric complexes consisting of both caveolin-1 and caveolin-2, the latter of which seems to be restricted to the basolateral membrane. Remarkably, caveolin-2 requires caveolin-1 for proper plasma membrane delivery because caveolin-2 is retained in the Golgi complex in caveolin-1-deficient Fischer rat thyroid cells or K562 erythroleukemia cells, but redistributes to the plasma membrane on coexpression of caveolin-1 (20, 28). Finally, epithelial cell lines devoid of caveolin-1 (Caco-2 or Fischer rat thyroid) do not contain caveolae, but reintroduction of caveolin-1 restores caveolar formation which again is restricted to the basolateral membrane (20, 49). Thus the overall conclusion for epithelial cells expressing caveolin-1 and -2 is that the apical membrane contains caveolin-1, but no caveolae, whereas the presence of both caveolin-1 and -2 in the basolateral membrane allows for the formation of caveolae (16). The polarized effect of caveolin-1 expression on caveolae formation is mirrored in our study by the selective stimulation of the hypotonicity-induced taurine and ATP efflux at the basolateral membrane of caveolin-1- expressing Caco-2 cells. This indicates that basolateral caveolin-1 and/or basolateral caveolae play a role either in the basolateral sorting of the transporters or in the formation of a functional signal transduction complex as discussed above (Fig. 7).
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In conclusion, with the use of stably transfected Caco-2 cells grown as monolayers, we show that caveolin-1 expression differentially affects the hypotonicity-induced release of taurine and ATP in a polarized epithelial cell by increasing the basolateral release. The present data in combination with our previous observations on VRAC stimulation by caveolin-1 (45) are consistent with a modulating/facilitating role for caveolin-1 in the functional expression of swelling-induced efflux of Cl–, organic osmolytes, and ATP. It remains to be investigated how caveolin-1 achieves this effect, e.g., by promoting plasma membrane delivery of ion channels or transporters or formation of a signal transduction complex.
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ACKNOWLEDGMENTS |
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Present address for N. Ullrich, Cardiac Medicine, National Heart and Lung Institute, Imperial College London, Dovehouse St., London, SW3 6LY, United Kingdom.
Present address for A. Caplanusi: Department of Medical Biochemistry, Carol Davila University of Medicine and Pharmacy, Eroii Sanitari Blvd. 8, R-050474 Bucharest, Romania.
Present address for B. Brône: Global Clinical Operations, Johnson&Johnson, Turnhoutseweg 30, 2340 Beerse, Belgium.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
* N. Ullrich and A. Caplanusi contributed equally to this work. 
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in A and B). E: pooled data (means ± SE) for the membrane current at +80 mV under ISO conditions or after stimulation with I/F and HTS in WT Caco-2 (n = 17) or caveolin-1-transfected Caco-2 (cav1-K; n = 39). *P < 0.05, statistically significant differences.
