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【摘要】 The epithelial sodium channel is found in apical membranes of a variety of native epithelial tissues, where it regulates sodium and fluid balance. In vivo, a number of hormones and other endogenous factors, including polyunsaturated fatty acids (PUFAs), regulate these channels. We tested the effects of essential n-3 and n-6 PUFAs on amiloride-sensitive sodium transport in A6 epithelial cells. Eicosapentaenoic acid [EPA; C20:5(n-3)] transiently stimulated amiloride-sensitive open-circuit current ( I Na ) from 4.0 ± 0.3 to 7.7 ± 0.3 µA/cm 2 within 30 min ( P < 0.001). No activation was seen in the presence of 10 µM amiloride. In cell-attached but not in cell-excised patches, EPA acutely increased the open probability of sodium channels from 0.45 ± 0.08 to 0.63 ± 0.10 ( P = 0.02, paired t -test). n-6 PUFAs, including linoleic acid (C18:2), eicosatetraynoic acid (C20:4), and docosapentanoic acid (C22:5) had no effect, whereas n-3 docosahexanoic acid (C22:6) activated amiloride-sensitive I Na in a manner similar to EPA. Activation of I Na by EPA was prevented by H-89, a PKA inhibitor. Similarly, PKA activity was stimulated by EPA. Nonspecific stimulation of phosphodiesterase activity by CoCl 2 completely prevented the effect of EPA on sodium transport. We conclude that n-3 PUFAs activate epithelial sodium channels downstream of cAMP in a cAMP-dependent pathway also involving PKA.
【关键词】 patch clamp epithelial sodium channel PKA A cells amiloride
-3 POLYUNSATURATED FATTY acids (n-3 PUFAs) have been implicated as important dietary factors in the prevention of heart disease in both epidemiological studies and clinical prevention trials ( 9 ). These essential fatty acids are found in fish and plants and are consumed as dietary supplements due to their putative protective role. The n-3 PUFAs have effects on a diversity of physiological processes, suggesting that they may play multiple roles in cardiovascular health including triglyceride and cholesterol metabolism as well as blood pressure regulation, thereby implicating various cellular mechanisms. Besides their effects on channel gene expression ( 21 ), PUFAs activate membrane enzymes, channels, and transporters by covalent modification of the proteins, alteration of their interaction with the membrane lipid bilayer, or by modification of the biophysical properties of the membrane itself ( 10, 19, 20, 29, 37 ).
Essential n-3 PUFAs also are modulators of ionic conductances, in particular of the cardiac voltage-gated sodium channel and of various voltage-gated and calcium-activated potassium channels, suggesting a direct effect on cardiac electrical activity ( 8, 20, 27 ). Little is known about the effects of n-3 PUFAs on other ion channels, particularly in epithelia.
The current study was undertaken to examine the effect of n-3 PUFAs on the epithelial renal cell line A6, which expresses highly selective sodium channels ( 14 ). These channels are essential in the physiological maintenance of sodium balance and are regulated by a number of hormones and endogenous factors, including aldosterone, insulin, antidiuretic hormone, and prostaglandins, which act through a variety of posttranslational modifications, including PKA and PKC phosphorylation, methylation, and ubiquitination ( 2, 7, 22, 25, 30, 35 ).
In this study, we observed a rapid and reversible stimulatory effect on transepithelial sodium transport by the n-3 PUFA eicosapentanoic acid [EPA; n-3(C20:5)]. This stimulation is due to increased apical membrane permeability to sodium. EPA activates the amiloride-inhibitable sodium channel protein in a cAMP-dependent pathway involving PKA. The increased level of sodium transport and modulation of the biophysical properties of epithelial sodium channels by -3 fatty acids point to the potential importance of these dietary essential fatty acids in maintaining sodium balance and regulating blood pressure.
EXPERIMENTAL PROCEDURES
Cell culture. A6 cells (American Type Culture Collection derived originally from Xenopus laevis, passages 74-82 ) were maintained in culture on plastic flasks in DMEM/F-12 growth medium (Invitrogen), adapted for amphibian tissue culture osmolarity by a 20% dilution with distilled water and supplemented with 5% FBS (HiClone), 25 U/ml penicillin, and 25 µg/ml streptomycin. Cells were grown at 28°C in 1% CO 2. For electrical measurements, cells were grown on permeable supports for 10 days (12-mm-diameter inserts, Transwell, Costar). Maximum and stable values of transepithelial sodium transport and electrical parameters are observed at this time, indicating that apical sodium channels are functional. For patch-clamp experiments, cells were plated on transparent polyester filters (Snapwell inserts, 12 mm in diameter, Transwell) and for biochemical work, on 100-cm 2 homemade structures with porous supports (HAWP, Millipore). All experiments were performed after 10 days of culture. Serum was omitted in the last 24 h.
Transepithelial voltage and resistance were measured using an EVOM volt-ohmmeter (World Precision Instruments). The corresponding amiloride-inhibitable sodium current ( I Na ) was calculated from these values. I Na is used as an estimate of the net transepithelial sodium current under the conditions of this study. All measurements were performed in parallel on control and treated tissues.
Patch-clamp experiments. Snapwell inserts were placed in the experimental chamber filled with solution, and cells were viewed using a Nikon Diaphot inverted microscope (Tokyo, Japan). Patch pipettes with resistances of 15-20 M were constructed using borosilicate glass capillaries (Hilgenberg, Malsfeld, Germany), pulled in two stages with a PB-7 puller (Narishige, Tokyo, Japan), and used without polishing. Single-channel currents were obtained at room temperature and amplified using an Axopatch-200A amplifier (Axon Instruments). Current records were low-pass filtered at 300 Hz through an eight-pole Bessel filter (900 LPF, Frequency Devices, Haverhill, MA) and acquired online at the rate of 1 kHz using a TL-1 DMA interface and Axotape 1.2 software (Axon Instruments).
Twenty-four hours before the experiment, the cell culture medium was switched to diluted DMEM/F-12 (120 mM Na, osmolarity 240 mosmol/kgH 2 O) without serum and antibiotics. Sodium bicarbonate was replaced by HEPES (10 mM, pH = 7.4). For cell-attached experiments, this medium was also used for the extracellular bath and patch pipette solutions. To ensure rapid, constant flow, cells were continuously superfused by gravity feed at the rate of 0.5 ml/min through Teflon tubing connected to a manifold, and outflow tubing was positioned next to the area of patch formation. EPA was added to the solution just before the experiment. Control experiments were run in the presence of ethanol, used to dissolve EPA.
Cell-attached configuration. Recording started when seal resistance of the patch exceeded 20 G ( 5 min). Baseline channel activity was recorded for 5 min before the apical solution was replaced with the solution containing EPA. The recordings were continued for as long as the integrity of the patch was maintained (usually up to 20 min).
Excised patches. Inside-out patches were bathed with a solution containing (in mM) 85 KCl, 3 NaCl, 4 CaCl 2, 1 MgCl 2, 5 EGTA, and 10 HEPES (pH 7.4 adjusted with 1 N KOH, osmolarity 235 mosmol/kgH 2 O).
The applied voltage in cell-attached and inside-out configurations results in a deflection from the patch potential (i.e., the resting membrane potential for cell-attached patches and 0 mV for inside-out excised patches). According to convention, inward sodium channel currents (pipette to cell) are represented as downward transitions in single channel records.
PKA activity. PKA activity was measured using a SignaTECT cAMP-dependent PKA assay kit according to the manufacturer's instructions (Promega). Briefly, this method involves measuring the transfer of 32 P-labeled phosphate to a specific PKA substrate (biotinylated Kemptide). Cells were washed twice with PBS, scraped in cold extraction buffer (25 mM Tris·HCl, pH 7.4; 0.5 mM EDTA, 0.5 mM EGTA, 10 mM -mercaptoethanol, 1 µg/ml leupeptin, 1 µg/ml aprotinin, 0.5 mM PMSF), homogenized with a Dounce homogenizer, and centrifuged at 14,000 g for 5 min at 4°C. The supernatants were kept on ice and assayed immediately.
Chemicals. Unless otherwise noted, all chemicals were purchased from Sigma-Aldrich.
Data analysis and statistics. Single-channel analysis was performed using pClamp (Axon Instruments) and WinASCD software (Laboratorium voor Fysiologie, Leuven, Belgium). Data records were low-pass filtered at 100 Hz. All-points histograms were obtained from current records by Fetchan software followed by their baseline correction using the WinASCD program. The single-channel amplitudes and open probability ( NP o ) were determined from all event lists of single-channel records. NP o, the product of the number of channels in a patch ( N ) by the open probability, which reflects channel activity within a patch, was calculated using the following equation
where T is the total recording time, i is the number of open channels, t i is the recording time during which i channels were open, and N is the apparent number of channels within the patch determined as the highest observable level. Therefore, NP o can be calculated without any assumptions about the total number of channels in a patch or the open probability of single channels.
All NP o values were calculated for 2.5-min intervals of recording and are reported as means ± SE. In previous patch-clamp studies in A6 cells from other groups, significant variability in baseline NP o is documented between individual patches ( 3, 23 ). For this reason, we conducted paired experiments, with each patch serving as its own control. Paired t -test analysis was performed for the average change in NP o, with a significance level of P < 0.05. A 2 -test was performed to compare the percentage of empty patches in control and experimental conditions (0.001, df = 1).
RESULTS
I Na. I Na was measured in cell monolayers after 10 days in culture. Serum was removed from the media 24 h before the experiment. EPA, whether added to the apical or to the basolateral side, induced an increase in current from 4.0 ± 0.3 to 7.7 ± 0.3 µA/cm 2 (means ± SE; n = 15 independent triplicates). This effect started after 5 min, reached a peak within 30 min, and returned to control values after 60 min ( Fig. 1 ). We did not observe any effect of EPA at concentrations below 33 µM, and higher concentrations did not further stimulate I Na (data not shown). The addition of 10 µM amiloride, a specific inhibitor of epithelial sodium channels, completely inhibited EPA stimulation of the current and, when added before EPA, prevented its effect (data not shown). Readdition of EPA after the return of the current to control values caused a second activation with similar magnitude and kinetics.
Fig. 1. Time course of stimulation of transepithelial sodium current ( I Na ) in A6 monolayers. Control and eicosapentaenoic acid (EPA)-treated tissues were measured in parallel. Control wells received an equivalent amount of ethanol, and control values were stable for the duration of experiments (not shown). Data are expressed as percent increase compared with time 0 values of experimental tissues. EPA, prepared in ethanol, was added to the apical bath at a final concentration of 33 µM (1:1,000 dilution). Measurements were carried out in triplicate ( n = 15). * P < 0.01. ** P < 0.001, repeated measures ANOVA.
Docosahexanoic acid (DHA; C22:6), another n-3 PUFA, activated amiloride-sensitive I Na similar to EPA, whereas n-6 PUFAs such as linoleic acid (C18:2), eicosatetraynoic acid (C20:4), or docosapentanoic acid (C22:5) had no effect on I Na (data not shown). Because the results obtained with the two n-3 PUFAs (DHA and EPA) were similar, only the EPA data will be shown throughout the paper.
Because increases in transepithelial sodium transport may be attributed to increases in basolateral Na + -K + ATPase pump activity or apical membrane sodium permeability, we first examined whether the stimulation of current by EPA resulted from a nonspecific effect on the basolateral pumps. To this end, apical sodium channels were completely inhibited by amiloride, but sodium was allowed to enter via pores induced by the ionophore nystatin in the apical solution. Pump activity was maintained below V max by using low sodium in the apical Ringer solution (12 mM). Under these conditions, EPA addition did not increase the current, indicating no direct effect on the basolateral Na + -K + -ATPase.
Patch-clamp measurements of apical sodium channels. We measured apical sodium single-channel activity in cell-attached patch-clamp experiments. Because the effect of EPA fully develops within about 20 min, we first checked the single-channel activity in untreated patches during this time interval. A representative record is shown in Fig. 2 A. The timescale magnification of current traces shows visible decay in channel activity, which was further confirmed by all-point histogram analysis ( Fig. 2 A, insets ). The corresponding time course of calculated NP o is shown in Fig. 3 A (open bars). Rundown of channel activity was observed after 10-15 min in seven of the nine control patches, with NP o decreasing from 0.50 ± 0.06 to 0.43 ± 0.05 ( P = 0.01, paired t -test). This decay in activity could obscure a stimulatory effect of EPA, especially as we chose to use initial NP o as the internal control for the effect of EPA (see EXPERIMENTAL PROCEDURES ). A similar decay was also reported by other investigators ( 3, 24 ). However, after the addition of EPA, single-channel activity increased. A representative record of three experiments is shown in Fig 2 B. Timescale magnification of current traces and all-point histogram analysis clearly indicate activation of channel activity. Like the observed transepithelial increase in current, the increase in channel activity started after 5 min and was observed for over 15 min, after which time the stability of the patches spontaneously deteriorated, preventing further observations. The corresponding time course is shown in Fig. 3 A (hatched bars). On average, NP o increased from 0.45 ± 0.08 to 0.63 ± 0.10 after addition of EPA ( P = 0.02, paired t -test).
Fig. 2. Single-channel activity in A6 cell-attached patches. A : representative record from a control patch under constant vehicle perfusion conditions. Timescale magnification of traces in selected intervals are shown. All-point histograms were constructed from 200-s periods at the begining and end of the record ( insets ). Currents were digitally filtered at 30 Hz. B : representative record from a patch in which vehicle perfusion was switched after 5 min to a solution containing 33 µM EPA. Timescale magnification of traces in selected intervals are shown. All-point histograms were constructed from the 200-s period preceding EPA addition and the last 200-s period of the record ( insets ). Currents were digitally filtered at 30 Hz.
Fig. 3. A : time course of open probability ( NP o ) for the current records shown in Fig. 2 A. The NP o values, calculated for each 2.5-min interval, are shown for control (open bars) and experimental recordings (hatched bars). Arrow indicates when perfusion of the experimental patch was switched to a solution containing 33 µM EPA. B : current-voltage relationships of the sodium channels from control patches (open symbols) and from patches pretreated with 33 µM EPA for 15-30 min (filled symbols). Data points represent means ± SE, and the lines are the linear fits to the data with a slope of 4.9 pS. - V p, pipette voltage. C : EPA does not activate sodium channels in cell-free inside-out patch. Single-channel records were obtained after excision of the patch under control conditions ( top trace) and after a 5-min perfusion with 33 µM EPA ( bottom trace). Bars indicate closed level (C), and downward deflections represent inward sodium current. Currents were digitally filtered at 50 Hz.
Exposure of cells to EPA for 15-30 min before patch formation had no significant effect on the apparent number of channels within the patch (6.2 ± 3.1 channels vs. control 4.9 ± 2.8, P 0.05) or on the number of patches without channel activity (10 of 23 vs. 83 of 182 for controls, P = 0.98, 2 -test). These results suggest that the effect of EPA is solely on the open probability. However, due to limitations in the calculation of the true number of channels in the patches ( 26 ), a small effect of EPA on N cannot be excluded.
Figure 3 B shows the current-voltage relationships for sodium channels from cell-attached patches. The slope of channel conductance within the physiological range of transmembrane potentials (-100 to +50 mV) did not change in the presence of EPA ( = 4.9 ± 0.5 pS). The actual apical membrane potential cannot be measured and controlled in the cell-attached configuration. Hence, the apparent parallel shift in the current-voltage curve most likely represents depolarization of the apical membrane due to increased sodium entry into the cells. It is thus clear that EPA does not stimulate transepithelial sodium transport through modification of the unit conductance or the ion selectivity of apical sodium channels.
We also studied single-channel activity in cell-free inside-out patches. Immediately after withdrawal of the pipette, channel activity rapidly decreased, approaching zero after 30 s in six of eight patches. This agrees with other studies of A6 cells ( 24, 40 ). In contrast to the results obtained in cell-attached patches, application of EPA neither increased the activity of remaining channels nor reactivated channels after rundown ( Fig. 3 C ). Together, this observation and the 5-min delay before any effect of EPA is observed suggest that EPA does not directly affect sodium channel activity in A6 cells.
Involvement of prostaglandins and leukotriens. Polyunsaturated fatty acids with 20 carbon atoms, such as EPA, are transformed into metabolically active eicosanoids including prostaglandins and leukotriens. To test whether EPA acted through these metabolites, I Na was measured in the presence of inhibitors of cyclooxygenase and of lipooxygenase. Inhibition of prostaglandin and leukotrien formation by indomethacin and nordihydroguariaretic acid, respectively, did not prevent stimulation of current by EPA ( Fig. 4 A ).
Fig. 4. I Na in A6 monolayers. A : inhibition of eicosanoid production. Control and treated tissues were measured in parallel. Data are expressed as percentage of time 0 value (first bar). The effect of EPA (33 µM) is shown at 30 min (second bar; ** P < 0.001, unpaired t -test, n = 3). Cells were incubated with nordihydroguariaretic acid (NDGA; 2 µM in ethanol) and indomethacine (2 µM in ethanol) for 1 h (third bar) before addition of 33 µM EPA (last bar). Values obtained with EPA plus inhibitors were not significantly different from EPA alone (unpaired t -test). B : PKA pathway. Data are expressed as percentate of the time 0 (first bar). Effect of 33 µM EPA is shown at 30 min (second bar). H-89, an inhibitor of PKA (5 µM in ethanol, third bar) was added 30 min before the addition of 33 µM EPA for 30 min. Stimulation of current by EPA (** P < 0.001) was significantly lowered by H-89 (last bar, # P < 0.01, n = 3). Values obtained with H89 or H89 plus EPA were not significantly (NS) different from control (unpaired t -test). C : cAMP requirement. Data are expressed as percentage of time 0 (first bar). Effect of 33 µM EPA is shown at 30 min (second bar). CoCl 2 (1 mM in water, third bar) was added 30 min before the addition of 33 µM EPA for 30 min (last bar). EPA stimulation of current (** P < 0.001) was significantly lowered by CoCl2 (last bar, # P < 0.008, n = 3). Values obtained with CoCl 2 or CoCl 2 +EPA were not significantly different from control (unpaired t -test).
PKA pathway. cAMP-dependent PKA phosphorylation plays little role in the basal rate of sodium transport ( 5 ) but, when stimulated by vasopressin or forskolin, leads to increased sodium reabsorption ( 26, 30 ). H-89, an inhibitor of PKA, had no effect on control I Na but prevented the stimulation by EPA ( Fig. 4 B ), suggesting the involvement of the cAMP-dependent PKA phosphorylation pathway in the action of EPA. Similarly, after the current was stimulated with forskolin, EPA did not further increase I Na (data not shown). Furthermore, nonspecific stimulation of PDE activity by CoCl 2, which leads to decreased levels of intracellular cyclic nucleotides, completely prevented the effect of EPA ( Fig. 4 C ). Additional evidence for PKA-dependent phosphorylation was obtained by direct PKA activity measurements. PKA activity increased by 66% after 15 min of apical application of EPA (33 µM). Under the same experimental conditions, 0.1 µM vasopressin, used as a positive control, stimulated PKA activity by 127% ( Fig. 5 ).
Fig. 5. Effect of EPA on PKA activity. Assay was performed on A6 cells cultured in serum-free media for 24 h. Fifteen minutes before the assay, cells received either vehicle (control), EPA (33 µM), or vasopressin [0.1 µM antidiuretic hormone (ADH) positive control]. Data are means ± SE of triplicate measurements. The difference between treated cells and control is statistically significant (unpaired t -test * P < 0.001).
DISCUSSION
In the present study, we combined transepithelial current measurements with single-channel recordings in the cultured cell line A6 commonly used as a model of the distal nephron for more than 20 years ( 14 ).
We found that n-3 PUFAs, and particularly DHA and EPA, exert a stimulatory effect on sodium transport that is specific (completely blocked by amiloride), rapid, transient, and dependent on cyclic nucleotide availability. This effect was not mediated by the metabolites of essential fatty acids, prostaglandins, and leukotriens.
Fatty acid regulation of epithelial sodium conductance has been described in fetal alveolar cells ( 1, 13 ), in A6 cells ( 4, 39 ), and in the oocyte expression system ( 4 ). The fatty acids investigated in those studies were eicosatetraynoic acid and arachidonic acid, which are both -6 fatty acids. These fatty acids increased single-channel epithelial sodium channel activity in fetal pneumocytes ( 1 ) but lowered both single-channel activity ( 39 ) and channel protein expression ( 4 ) in A6 cells and the oocyte expression system.
A number of regulatory pathways control sodium channel activity directly or indirectly in native epithelia, including methylation and a number of phosphorylation (mediated by either PKC, PKA, SGK1, Src kinase, MEK, or p38 MAPK) and dephosphorylation reactions ( 2, 6, 7, 15, 17, 18, 23, 25, 30, 32, 35, 36 ).
Further insight into the mechanism of sodium channel activation by EPA was gained from results obtained with CoCl 2, H-89, and direct measurements of PKA activity, which showed that cyclic nucleotides and PKA activity were required for the effect of EPA. The transepithelial measurements suggested that EPA brings sodium transport to maximal capacity because there was no further increase when EPA was re-added at the peak of the first stimulation.
The substantial number of empty patches found in both control and EPA-treated cells suggests a heterogeneous distribution of channels, which may reflect the previously described compartmentalization of active sodium channels within membrane microdomains ( 16, 33 ). The effect of EPA on active patches was an increased NP o of the channels. Increases in NP o can result from cAMP-dependent PKA phosphorylation of channels ( 11, 26 ), cAMP-mediated insertion of new channels ( 26, 34 ), or an increase in the turnover rate by ubiquitin conjugation of the channels ( 31 ), none of the which is mutually exclusive.
The increased sodium transport observed with EPA in renal epithelium leads us to extrapolate that the reported drop in blood pressure observed with EPA both in hypertensive rats ( 12, 38 ) and in patients suffering from essential hypertension ( 28 ) could be related to the attenuation of the renin-angiotensin-aldosterone axis, secondary to the increased renal sodium reabsorption. The blood and cellular concentration of EPA depends on its intake in the diet, its synthesis from -linolenic acid, and its further metabolism. The total EPA pool in plasma can be estimated from phospholipids, triglycerides, cholesteryl esters, and free fatty acid fractions. Assuming that EPA represents 0.3-1% of each of these fractions, biologically relevant concentrations of EPA 33 µM would be expected, and these would increase several-fold after fish oil feeding,
GRANTS
This research was supported by funds from the Université Libre de Bruxelles and Fonds Defay. V. Shlyonsky is the recipient of a postdoctoral fellowship from the Belgian National Research Foundation (FNRS), and A. Goolaerts is a doctoral fellow from the FNRS.
ACKNOWLEDGMENTS
We thank Profs. R Naeije and P. Calder for help and support and N. Mason for the preliminary I Na measurements with fatty acids.
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作者单位：Physiology Department, Université Libre de Bruxelles, 1070 Brussels, Belgium