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A variety of studies have shown that Na+ reabsorption across epithelial cells depends on the protease–antiprotease balance. Herein, we investigate the mechanisms by which α1-antitrypsin (A1AT), a major anti-serine protease in human plasma and lung epithelial fluid and lacking a Kunitz domain, regulates amiloride-sensitive epithelial Na+ channel (ENaC) function in vitro and in vivo. A1AT (0.05 mg/ml = 1 μM) decreased ENaC currents across Xenopus laevis oocytes injected with human α,β,γ-ENaC (hENaC) cRNAs, and human lung Clara-like (H441) cells expressing native ENaC, in a partially irreversible fashion. A1AT also decreased ENaC single-channel activity when added in the pipette but not in the bath solutions of ENaC-expressing oocytes patched in the cell-attached mode. Incubation of A1AT with peroxynitrite (ONOO−), an oxidizing and nitrating agent, abolished its antiprotease activity and significantly decreased its ability to inhibit ENaC. Intratracheal instillation of normal but not ONOO−-treated A1AT (1 μM) in C57BL/6 mice also decreased Na+-dependent alveolar fluid clearance to the same level as amiloride. Incubation of either H441 cells or ENaC-expressing oocytes with normal but not ONOO−-treated A1AT decreased their ability to cleave a substrate of serine proteases. A1AT had no effect on amiloride-sensitive currents of oocytes injected with hENaC bearing Liddle mutations, presumably because these channels remain at the surface longer than the wild-type channels. These data indicate that A1AT may be an important modulator of ENaC activity and of Na+-dependent fluid clearance across the distal lung epithelium in vivo by decreasing endogenous protease activity needed to activate silent ENaC.
Our data show that α1-antitrypsin (A1AT) inhibits active Na+ reabsorption and lung fluid clearance across distal lung epithelial cells and highlight a potential mechanism by which A1AT administration may prove beneficial in clinical situations in which increased ENaC activity may contribute to lung pathology (such as cystic fibrosis).
The amiloride-sensitive epithelial Na+ channels (ENaCs) play an important role in Na+ and fluid homeostasis across a number of epithelial cells, including those in airway and alveolar spaces (1, 2). ENaC mRNA, protein levels, and ENaC activity are regulated by a variety of hormones (such as aldosterone and dexamethasone), second messengers cAMP/protein kinase A, and reactive intermediates (2–4). In addition, endogenous channel-activating proteases (CAPs), as well as proteases released by inflammatory cells (trypsin, elastase), activate ENaC either by cleaving critical amino acids in α- and γ-ENaC subunits, or by activating signaling pathways (1, 2, 5–7).
There has been considerable interest in identifying the mechanisms by which endogenous and exogenous proteases activate ENaC. Aprotinin, a potent and reversible Kunitz-type inhibitor of several serine proteases, including trypsin, plasmin, and kallikreins (8), has been reported to inhibit sodium transport among a variety of epithelial cells, including A6 (9), human bronchial epithelial (HBE) cells (10, 11), and rat and mouse lung alveolar epithelial cells (1). Other Kunitz-type serine protease inhibitors, such as hepatocyte growth factor activator inhibitor (HAI)-1 and HAI-2 (placental bikunin), have also been demonstrated to inhibit prostasin and ENaC activity (12). Thus, it is generally accepted that Kunitz-type protease inhibitors block epithelial serine proteases needed for ENaC activation, and these inhibitors have been suggested as a novel treatment for the Na+ hyperabsorption in cystic fibrosis (CF) airways (10).
α1-Antitrypsin (A1AT) is an acute-phase glycoprotein and a member of the serine protease inhibitor (SERPIN) superfamily (13). Until recently, it was thought that its primary function was to neutralize elastase in vivo, protease-3, and other serine proteases released from activated human neutrophils. In fact, the rate of formation of the A1AT/neutrophil elastase inhibitory complex is one of the fastest known for SERPINs (6.5 × 107 M−1 s−1). However, recent findings indicate that A1AT also directly inhibits the activity of caspase-3, a cysteine protease that plays an essential role in apoptosis (14, 15), and the catalytic domain of recombinant human matriptase-1 (16), a multidomain type-II transmembrane serine protease, which is involved in ENaC proteolysis and activation (17). These findings lead to the hypothesis that both Kunitz-type and non–Kunitz-type serine protease inhibitors may inactivate epithelial serine proteases, and thus prevent proteolytic activation of ENaC in vivo.
Herein we performed a series of biophysical, biochemical, and physiological studies in Xenopus oocytes, injected with human α,β,γ-ENaC (hENaC) cRNAs, human lung Clara-like cells (H441), which express ENaC type-single channel activity (18), as well as in anesthetized mice, which clear alveolar fluid across their distal epithelium secondary to active Na+ transport (19, 20), to identify the mechanisms by which A1AT inhibits amiloride-sensitive Na+ transport and alveolar fluid clearance (AFC) both in vitro and in vivo. Our results indicate that very low concentrations (0.05 mg/ml or 1 μM) of A1AT decreased amiloride-sensitive currents (Iamil) in oocytes and H441 cells in a partially irreversible fashion, and reduced Na+-dependent AFC in vivo. Additional studies showed that these effects were the result of A1AT inhibition of membrane-bound proteases present on the surface of oocytes and H441 cells. These new findings indicate that the inhibitory effect of A1AT might be an important regulator of ENaC activity and Na+-dependent AFC in vivo.
Purified human A1AT was purchased from Calbiochem (La Jolla, CA) and Sigma-Aldrich (St. Louis, MO). Prolastin was provided by Talecris Biotherapeutics, Inc. (Research Triangle Park, NC). The purity of A1AT preparations was greater than 97%, and their inhibitory activity was higher than 75%. Native A1AT was diluted in PBS (pH 7.4). To ensure the removal of endotoxin, A1AT was eluded through Detoxi-Gel AffinityPak columns according to manufacturer instructions (Pierce, Rockford, IL). Purified batches of A1AT were then tested for endotoxin with the limulus amebocyte lysate endochrome kit (Endosafe; Charles River, Charleston, SC). Endotoxin levels were less than 0.1 enzyme units/mg protein in all preparations used.
A1AT (1.25 mg/ml) was suspended in PBS containing 10 mM HEPES (pH 7.4) and incubated with peroxynitrite (ONOO−; 1 mM). Because ONOO− has a half-life of less than 1 second at pH 7.4 (21), it was added while still in NaOH as the solution was vigorously mixed. The pH was carefully monitored to ensure that reactions were performed at neutral pH. A1AT was then separated by SDS-PAGE (8 or 12% separating gel) in the presence of 2-mercaptoethanol, and visualized by staining the gels with GelCode (Thermo Fisher Scientific, Rockford, IL).
α,β,γ-hENaC cDNAs with the Liddle mutations (hENaC-α595x, hENaC-β566x, hENaC-βS520K, hENaC-γ575x in a pMT3 vector) were a kind gift from Dr. Peter Snyder (University of Iowa). These plasmids were designed for nuclear injection. ENaC subunits were subcloned into pCDNA3.1 vector for in vitro transcription. hENaC-α595x was excised by NotI and Xho and cloned into pCDNA3.1 at NotI and XhoI. hENaC-β566x and βS520K were excised by NotI and Acc65I and cloned into pCDNA3 at NotI and XhoI, with Acc65I and XhoI blunt ended. hENaC-γ575x was excised by NotI and EcoRI and cloned into pCDNA3.1 at NotI and XhoI, with EcoRI and XhoI blunt ended. All the constructs were verified by DNA sequencing. ENaC subunit plasmids were linearized and cRNAs were prepared with a cRNA synthesis kit (Message Machine T7; Ambion, Austin, TX) according to the manufacturer's protocol. cRNAs were dissolved in RNAse-free water, and the concentrations were determined spectrophotometrically.
Detailed description of these techniques has been previously reported (22–24). In brief, defolliculated oocytes, isolated from Xenopus laevis frogs, were injected with cRNAs encoding for wild-type α, β, γ-hENaC (8.4 ng each), dissolved in 50 nl of RNase-free water per oocyte, and incubated in half-strength L-15 medium for 24 to 48 hours. Whole-cell cation currents were measured by the two-electrode voltage clamp technique before and after perfusion with amiloride (10 μM), which inhibited more than 90% of the inward Na+ currents. The oocytes were held in a small groove in an experimental chamber of 1-ml volume at room temperature (21°C). The chamber was filled with ND96 solution containing: 96 mM NaCl, 2.7 mM KCl, 1 mM MgCl2, 2 mM CaCl2, and 10 mM HEPES at pH 7.6 (osmolarity, 200–220 mosM). Oocytes were impaled with two 3 M KCl–filled electrodes, with resistances of 0.5–1.5 MΩ. A TEV 200 voltage clamp amplifier (Dagan Corp., Minneapolis, MN) was used to voltage clamp oocytes. Two reference electrodes were connected to the bath. The membrane potential (Vm) was held at −40 mV. Current–voltage (I–V) relationships were obtained by stepping the Vm from −120 mV to +80 mV in 20-mV increments for 600 milliseconds. Sampling protocols were generated by pCLAMP 8.0 (Axon Instruments, Molecular Devices, Union City, CA). Currents were sampled at the rate of 1 kHz, filtered at 500 Hz, and simultaneously stored electronically and displayed in real time on a chart recorder.
Two different sets of experiments were performed: in the first set, we measured I–V relationships before and after addition of amiloride (10 μM) into the bath solution containing ND96. Na+ currents were continuously recorded while stepping the Vm from −40 to −140 mV (to record inward Na+ currents) and from −40 to +60 mV (to record outward currents). Oocytes were then perfused with ND96 until stable currents were obtained at which point native or ONOO−-treated A1AT (0.05 mg/ml; 1 μM) were added into the bath solution and currents were continuously recorded until new steady-state values were reached (usually within 45 min). At that time, amiloride (10 μM) or trypsin (1 μM) was added to the bath and currents were recorded until new steady-states were reached. In a second set of experiments, hENaC-expressing oocytes were incubated for 2 hours in half-strength L-15 medium with and without A1AT (0.05 mg/ml; 1 μM). At the end of this period, they were placed in the recording chamber under the microscope (filled with ND96) and I–V relationships were recorded as described above.
In some cases, oocytes microinjected with hENaC were incubated with MG-132 (a proteasome inhibitor; Calbiochem) for either 24 hours (100 nM) or 2 hours (4 μM). After completion of I–V curves, they were perfused with A1AT (1 μM) and inward currents were recorded as described above.
Xenopus oocytes injected with either H2O, or hENaC cRNAs were incubated with either vehicle or A1AT (1 μM) for 45 minutes and then homogenized in 1 ml of 100 mM PBS (pH 7.4) containing one tablet of Complete Mini EDTA-free Protease Inhibitor Cocktail Tablets (Roche Applied Sciences, Indianapolis, IN) and 10 mM N-ethylmaleimide (NEM). Lysates were centrifuged twice at 300 × g for 10 minutes, and the supernatants collected and centrifuged at 16,000 × g for 30 minutes. The upper layer, consisting of lipid and yolk, was removed, and the remaining pellet and supernatants were suspended in 1% triton X100 and solubilized for 1 hour at 4°C. After centrifugation at 16,000 × g for 30 minutes, 300 μl of the extract was suspended in 1 ml of the same buffer, and γ-ENaC was immunoprecipitated with 5 μl of a polyclonal anti–γ-ENaC rabbit antibody (Abcam, Cambridge, MA) overnight at 4°C followed by 50 μl of 50% protein G beads for 3 hours at 4°C. The beads were washed extensively with the same buffer, followed by a final wash in 50 mM Tris (pH 7.5). Immune complexes were dissociated by heating in 60 μl of sample buffer at 95°C for 5 minutes, and probed with a rabbit polyclonal anti–γ-ENaC antibody (1:1,000 dilution; Abcam) followed by a secondary antibody (horseradish peroxidase protein A; 1:10,000 dilution; Pierce) and detected by chemiluminescence.
Surface expression of γ-hENaC was examined by a cell surface biotinylation assay, as previously described by us (25). cRNAs were coinjected into Xenopus oocytes. After 48 hours, oocytes were mechanically stripped of their vitelline membranes in hypertonic medium, and surface proteins were labeled with membrane-impermeable sulfo-NHS-Biotin (Pierce). Oocytes (10 per group) were subsequently lysed and centrifuged at 13,000 × g for 15 minutes at 4°C. Biotinylated proteins were precipitated with streptavidin–agarose (Pierce), and subjected to SDS-PAGE, followed by Western blotting with anti–γ-ENaC antibodies, as described above.
The vitelline membranes of oocytes expressing α,β,γ-hENaC were surgically removed after shrinking the oocytes by immersing them in a hypertonic sodium aspartate solution (400 mM). Immediately after removal of this membrane, oocytes were transferred to the recording chamber containing 0.5 ml ND96. Single channels were recorded in cell-attached mode using an Axopatch 200B amplifier (Axon Instruments). Signals were digitized at 5 KHz (DigiData 1,322 A, 16 bit; Molecular Devices, Sunnyvale, CA), filtered at 1 KHz, and acquired using an IBM compatible computer using the Axon pCLAMP 9.2 software (Axon Instruments).
Pipettes were made from a thick-walled capillary glass (LG 16; Dagan) using a two-stage puller (PC-10; Narishige, Tokyo, Japan). Pipette resistance was 9.7 (±0.9 MΩ) when filled with ND 96. The ground electrode (Ag-AgCl) was connected to the bath using a 3% agar bridge (1 M KCl). Single channels were recorded at a Vm of −100 mV. The mean value of the Vm, measured in a different set of oocytes using the double voltage clamp technique (18), was around 5.0 mV.
Open probability (Po) was measured as the total open time of a channel normalized to the total time of recording at −100 mV. For recordings showing more than one active channel (as was the case in most of our recordings), the average channels activity was reported as NPo (number of active channels multiplied by their Po), which is determined by integrating the area under the Gaussian curves to the all-points current–amplitude, and normalized to the peak area of the closed level or baseline current level. This routine is included in Clampfit software (Axon Instruments).
Clara-like H441 human lung cells were cultured on coverslips, as previously described (18). We added dexamethasone (100 nM) into the medium to enhance expression of ENaC and amiloride-sensitive Na+ currents. Whole-cell currents were elicited by applying a step-pulse protocol from −140 to +60 mV in 10-mV increments for 500 milliseconds from a holding potential of −40 mV. I–V relationships were constructed by averaging the current values between 400 and 500 milliseconds from the start of recording with the Clampfit program and plotted using Origin Software (Northhampton, MA). Only cells with stable baseline currents were included in the analysis. After obtaining stable whole-cell currents, cells were perfused with an external solution containing amiloride (2 μM) or an equivalent amount of H2O as vehicle, and amiloride-sensitive I–V relationships were calculated as described above.
Serine protease activity on the oocyte membrane and H441 cells were monitored by measuring the release of a fluorescent 7-amido-4-methyl coumarin (AMC) from a peptide substrate, t-Butyl oxy-carbonyl (Boc)-Gln-Ala-Arg-AMCHCl (R&D Systems, Minneapolis, MN) using a fluorescence plate reader (Fluoster Optima; BMG; Labtech GmbH, Durham, NC). Briefly, 10 oocytes were added to a cuvette containing 200 μM of ND96 medium, and the reaction was started by adding Boc-Gln-Ala-Arg-AMCHCl (50 μM final concentration). The reaction was monitored for up to 120 minutes by recording fluorescence at 460 nm after excitation at 380 nm. In parallel experiments, oocytes were pretreated for 45 minutes with A1AT (1 μM; either native or ONOO− treated) before recording fluorescence. The same protocol was followed with H441 cells, except that the cell number was adjusted to 105/ml in Ringer's solution. Finally, in some cases, measurements were obtained after disrupting the cellular membranes of either oocytes or H441 cells treated with 1 μM A1AT for 45 minutes.
AFC was measured as previously described (19, 20, 26). Briefly, C57BL/6 mice with body weights between 20 and 25 g were anesthetized with diazepam (1.0 mg/100 g, intraperitoneally; Abbott Laboratories, Abbott Park, IL) and ketamine (20 mg/100 g, intraperitoneal), and placed on a heating pad (Braintree Scientific, Cambridge, MA) to maintain body temperature at 37°C. The trachea was exposed and cannulated with a trimmed 18-gauge intravenous catheter, which was then connected to a mouse respirator (model 687; Harvard A1 Apparatus, Holliston, MA). Mice were paralyzed with pancuronium bromide (0.04 mg, intraperitoneal; Gensia Pharmaceuticals, Irvine, CA) and ventilated with 100% O2 with a 200-μl tidal volume (8–10 ml/kg body wt) at 160 breaths/min. Once stable anesthesia was achieved, mice were positioned in the left decubitus position, and 300 μl of isosmolar NaCl containing 5% fatty acid–free BSA was instilled via the tracheal cannula, followed by 100 μl of room air to clear dead space. In experimental groups, normal or ONOO−-treated A1AT (1 μM final concentration) was added into the instillate. Postinstillation, mice were ventilated for a 30-minute period, then the alveolar fluid was aspirated. AFC was calculated from the ratio of the protein concentration of the instillate before instillation and of the alveolar sample at 30 minutes.
All values are expressed as means (±1 SEM), along with the number of measurements. Iamil values were calculated from the difference between steady-state values before and after addition of amiloride (10 μM) into the bath. Data were analyzed by one-way ANOVA with Bonferroni's test or Student's paired t test, as appropriate. P values of less than 0.05 were considered significant.
In the first set of experiments, we assessed the extent and time course of inhibition of Iamil values by A1AT. Uninjected oocytes showed near-zero levels of inward currents. Thus, inward currents of ENaC-injected oocytes were attributed to the movement of Na+ ions, and will be referred to as “INa.” At a Vm of −100 mV, INa currents 48 hours after injection were −7,368 (±705; n = 171) (X ± 1 SEM; n = number of oocytes). Addition of amiloride (10 μM) into the bath inhibited more than 95% of INa. Addition of A1AT (obtained either from Calbiochem or Sigma) in the bath at a final concentration of 0.05 mg/ml (1 μM) inhibited INa by more than 90% within 45 minutes (Figures 1A and 1B). The mean rate constant (τ) of decay, calculated by fitting the INa with a monoexponential decay equation, y = A × exp(−INa/t) + B, was 13 (±0.6) minutes (X ± 1 SEM; n = 13). Adebamiro and colleagues (27) reported that aprotinin inhibited INa across A6 cells with a τ of 18 (±1.2) minutes. The INa inhibition by A1AT is most likely due to a decrease in the number of active ENaC channels at the plasma membrane.
Iamil values increased gradually after replacing the oocyte bath solution with ND96 medium without A1AT, reaching values of approximately 40% of controls by 6 hours (Figure 1C). Addition of trypsin (1 μM) into the bath after 45 minutes of incubation with A1AT restored INa to its control value within minutes (Figure 1B). Because trypsin is known to activate silent ENaC channels present in the cell membrane (28), our data suggest that A1AT decreases INa by inhibiting proteases that are needed for ENaC activation at the cell membrane. Both Prolastin (1 μM) and aprotinin (5 μM) inhibited INa to the same extent as A1AT (data not shown).
Exposure of A1AT to ONOO− oxidizes and nitrates a number of amino acids, including a key methionine (Met 358), located in the A1AT active site. These modifications were associated with loss of its ability to inactivate elastase (29, 30). SDS-PAGE analysis of normal A1AT revealed a single band at 52 kD, indicating that our preparations were free of contaminants from other antiproteases (Figure 2). On the other hand, higher molecular weight bands (in addition to the 52-kD band) were seen in SDS-PAGE blots of ONOO−-treated A1AT, consistent with self-aggregation (Figure 2). Western blotting studies of ONOO− (1 mM)-treated A1AT with a monoclonal anti-nitrotyrosine antibody and an alkaline phosphatase–conjugated monoclonal anti-dinitrophenyl group antibody (IgE) confirmed the presence of extensive nitration and oxidation (data not shown), in addition to the aggregation. In contrast to native A1AT, perfusion of ENaC-expressing oocytes with ONOO−-treated A1AT decreased INa by less than 15% within 45 minutes (Figures 1B). Similarly, incubation of ENaC-expressing oocytes with ONOO−-treated A1AT for 2 hours decreased INa by less than 15% (Figure 3). These data indicate that intact antiprotease activity is necessary for the A1AT inhibition of ENaC.
Recordings of single-channel activity in cell-attached patches of oocytes injected with α, β, and γ hENaC showed the presence of 4 picoSiemens (pS) channels with long open and closing times, characteristic of ENaC (Figure 4A). No change in channel activity (as assessed by NPo, the product of number of channels multiplied by their open probability) of cell-attached patches was seen for up to 15–20 minutes after A1AT addition into the bath. However, after 30 minutes, NPo increased from 0.15 (±0.014; n = 6) (control) to 3.02 (±0.52; n = 6) (A1AT; X ± 1 SEM; n = number of oocytes; P = 0.0003) (Figure 4B). The increase in NPo was most likely the result of decreased intracellular Na+ (due to the A1AT inhibition of ENaC), which, in turn, stimulated silent channels across the cell-attached patch, which was not exposed to A1AT (31). In contrast, addition of A1AT into the pipette decreased NPo from the control value of 0.15 (±0.014) to 0.077 (±0.019) (X ± SEM; n = 6; P < 0.01) (Figure 4B). These data indicate that the inhibitory effects of A1AT on ENaC require that A1AT must be present in its immediate vicinity.
Results shown so far indicate that A1AT inhibits amiloride-sensitive currents across oocytes expressing human ENaC. To assess the relevance of these findings in human lung epithelial cells, we patched H441 cells at the whole-cell mode and measured sodium currents (INa) before and after perfusion with 1 μM A1AT for 45 minutes. As shown in Figure 5, A1AT decreased H441 INa to the same extent as in Xenopus oocytes (Figure 1). Furthermore, trypsin restored a large fraction of the INa (Figure 5B). These results indicate that A1AT inhibits the activity of both heterologously expressed and native ENaC channels.
In the next series of experiments, we tested the possibility that the A1AT inhibition of ENaC was due to inactivation of endogenous proteases in Xenopus oocytes needed to cleave, and thus activate, ENaC. We preincubated ENaC-expressing oocytes and H441 cells with A1AT for 45 minutes, then added the fluorogenic peptide, Boc-Gln-Ala-Arg-AMC–HCl, which is cleaved by a number of serine and cysteine proteases, and measured fluorescence levels during the next 2 hours. As seen in Figure 6, there was a time-dependent increase of fluorescence levels when either oocytes or H441 cells were incubated with Boc-Gln-Ala-Arg-AMC–HCl, indicating the presence of active membrane-bound serine and cysteine proteases. Significantly lower fluorescence levels were seen when oocytes or H441 cells were preincubated with 1 μM A1AT. Interestingly, this decrease in fluorescence was reversed when oocytes and H441 cell membranes were ruptured by vigorous mixing while continuously measuring the fluorescence. This indicates the presence of intracellular proteases that were not inactivated by A1AT. Finally, normal levels of fluorescence were observed when either oocytes or H441 cells were preincubated with ONOO−-treated A1AT. These data indicate that normal but not ONOO−-treated A1AT inactivated membrane-bound serine proteases.
To further test the hypothesis that A1AT decreases ENaC activity by inactivating surface proteases necessary for its cleavage and activation, we tested the effects of A1AT on amiloride-sensitive currents of Xenopus oocytes injected with Liddle cRNAs. Liddle mutants have reduced binding for Nedd4-2 (32), a critical step leading to ubiquitination, internalization, and destruction by the proteasome and lysosome systems (32). Oocytes microinjected with hENaC cRNAs with the Liddle mutations, expressed higher levels of Iamil values 48 hours after injection as compared with those injected with wild-type hENaC (Figure 7A; −9991.908 ± 823.667 nA versus −6852.675 ± 727.541 nA for Liddle versus wild-type hENaC; n = 8; P = 0.01). A1AT had no effect on INa of these oocytes (Figure 6A), in sharp contrast to what was observed in oocytes injected with wild-type hENaC cRNAs (Figure 1). Based on these findings, we considered the possibility that A1AT decreased wild-type ENaC by promoting ubiqitination. We were unable to detect increased levels of ubiquitinated ENaC after treatment of Xenopus oocytes with A1AT (data not shown). Furthermore, incubation of oocytes expressing wild-type ENaC with a well known proteasome inhibitor (MG-132; 100 nM for 24 hours or 4 μM for 2 hours) delayed but did not alter the extent of A1AT inhibition of INa (Figure 7B). These findings argue against the possibility that A1AT enhanced ENaC ubiquitination. Instead, they suggest that Liddle ENaCs, because of their long residence time at the surface, were already cleaved before the incubation with A1AT, and did not require additional processing.
Incubation of ENaC-expressing oocytes with A1AT did not alter total levels of γ-ENaC; however, 20 minutes after incubation, at a time when INa was decreased by at least 50%, surface levels of cleaved, mature γ-ENaC (75-kD band), assessed by Western blotting of biotinylated (surface) proteins, were decreased considerably (Figures 8A–8C). Both in vitro and in vivo studies have shown that γ-ENaC at the plasma membrane is expressed in two distinct pools: the mature, active channels (~ 75-kD band; Figure 9B) that had undergone proteolytic processing, and immature, noncleaved channels (~ 93-kD band, Figure 9B), thought to be a reserve pool that can be activated by extracellular serine proteases (6, 33–35). Our results show that A1AT treatment decreased significantly the proteolytic cleavage and activation of surface γ-ENaC.
We and others have shown that anesthetized, ventilated mice clear a large fraction of intratracheally instilled fluid secondary to the active reabsorption of Na+ ions (20, 36–38). The first step in the vectorial transport of Na+ ions is their entry into the cytoplasm of alveolar epithelial cells through amiloride-sensitive channels down their electrochemical gradients (39, 40). As shown in Figure 9, AFC in normal C57BL/6 mice was approximately 30% of instillate per 30 minutes, and amiloride inhibited 50% of this value, in agreement with our previous findings in both C57BL/6 and BALB/c mice (20, 37). Addition of A1AT into the instilled fluid completely inhibited AFC to the same extent as amiloride. On the other hand, A1AT exposed to 1 mM ONOO− had no effect on AFC (Figure 9).
We report, for the first time, that native but not ONOO−-treated A1AT (which lacks antiproteolytic activity) down-regulates ENaC activity in both Xenopus oocytes expressing α, β, γ hENaC and in a human airway cell line (H441) expressing both Na+-selective channels with conductance of roughly 4 pS (characteristic of ENaC) and nonselective 20 pS cation channels (18). In addition, we show that intratracheal instillation of native but not ONOO−-treated A1AT in anesthetized mice decrease amiloride-sensitive Na+-dependent AFC across their distal lung epithelium. Only 40% of the Iamil values being restored 6 hours after A1AT washout from the bathing medium indicates that a fraction of the inhibition was irreversible at least within this time frame. A1AT plasma levels in normal individuals are approximately 1.3 mg/ml, and only 5–10% of plasma A1AT levels are found in the interstitial fluid and in the alveolar space (41, 42). Thus, the concentration of A1AT used both in our in vivo and in vitro studies (0.05 mg/ml = 1 μM) is close to its value in the mammalian epithelial lining fluid.
The number of active ENaC channels present at the cell membrane is regulated by two major mechanisms: (1) by proteolysis, which converts inactive channels to active channels; and (2) by removal of ENaC from the surface. The former requires the presence of membrane-bound serine proteases; the latter involves ubiquitination, internalization, and degradation of surface ENaC (2). We found that A1AT does not inhibit amiloride-sensitive currents in Xenopus oocytes expressing ENaC bearing Liddle mutations. We were unable to detect increased levels of ubiquitinated ENaC after treatment of Xenopus oocytes with A1AT, and, furthermore, incubation of oocytes expressing wild-type ENaC with proteasome inhibitor (MG-132) delayed, but did not alter, the extent of A1AT inhibition of INa. Taken together, our results allow us to conclude that A1AT regulates ENaC activity via the inhibition of the CAPs.
CAPs are membrane-anchored proteins with extracellular serine protease domains (9). Matriptase (CAP3) is an autoactivating protease expressed in most epithelia, and plays a major role in the activation of prostasin (CAP1), the protease that is considered responsible for γ-ENaC cleavage and maturation at the cell surface, in addition to the role played by furin (1, 5, 9). CAP1, CAP2, and CAP3 mRNAs, as well as CAP1 protein, have been shown to be present in mouse alveolar epithelial cells (1). Herein, we demonstrate the presence of such proteases on cell surface of oocytes and H441 cells using a specific substrate that releases a fluorescent probe (AMC) indicative of its cleavage by a serine protease at an arginine residue, similar to the site of ENaC subunits cleavage. Our data did not allow us to definitively identify which proteases were inhibited by A1AT. To the best of our knowledge, A1AT inhibits neither prostasin nor furin. Because prostasin is synthesized as an inactive zymogen, requiring activation by an upstream protease (matriptase) (43), we propose that one mechanism by which A1AT inhibits ENaC activity is by inactivating matriptase. In support of this hypothesis, recently, Tseng and colleagues (44) isolated A1AT/matriptase complexes from human milk that are resistant to dissociation in boiling SDS, suggesting that A1AT may bind to matriptase in an irreversible fashion. We also reported that native but not the inactivated A1AT inhibits the catalytic domain of matriptase with a second-order rate constant of 0.31 × 103 M−1s−1 in vitro in an irreversible fashion (16). Other serine proteinase inhibitors, like aprotinin and nexin-1 (PN-1) have also been shown to inhibit the activity of matriptase, thereby decreasing ENaC currents.
It is important to point out that, previously, A1AT (a member of the SERPIN family, which lacks the Kunitz-type domain) was found to be ineffective in inhibiting Na+ short-circuit currents across HBE cells mounted in Ussing chambers (10). A1AT (which does not inactivate furin) had also no effect on Na+ equivalent short-circuit current of mouse cortical collecting duct cells (34). On the other hand, a variant of A1AT (α1-PDX), which inhibits furin, significantly decreased the equivalent short-circuit current of cortical collecting duct (CCD) cells (34). In contrast, our data show that A1AT down-regulates both Iamil values in vitro and Na+-dependent AFC across the distal lung epithelia of anesthetized mice in vivo. Recently, Myerburg and colleagues (45) reported that protease inhibitor, PN-1, another member of the SERPIN family, inhibited Na+ currents across human airway epithelial cells isolated from CF lungs (45). Protease inhibitor PN-1, like A1AT, inhibits matriptase (45). Thus protease inhibitors with or without a Kunitz domain may play an important role in the protease–antiprotease balance responsible for ENaC processing and activation. Currently, the reasons for the different effects of A1AT on ENaC activity among these studies and ours are not clear. It is possible that ENaC cleavage in HBE and CCD cells was independent of matriptase or another protease inhibited by A1AT.
Aprotinin inhibition of airway cell INa was reversed either by removal of aprotinin or by addition of excess exogenous proteases, such as trypsin and elastase (10). Trypsin and elastase are serine proteases, known to activate ENaC channels in oocytes, human bronchial cell lines, and rat alveolar epithelial cells (9, 46), either by activating near-silent channels that are already present in the membrane (28), or by activating signal transduction pathways (7, 47). In our experimental model, the effect of A1AT on ENaC in oocytes was not immediately reversible; only 40% of the initial current was restored 6 hours after A1AT washout from the bathing medium, indicating that a fraction of the inhibition was irreversible at least within this time frame. One possible explanation can be that A1AT builds a stable complex with membrane-bound proteases (such as matriptase), and, therefore, part of A1AT remains on the membrane after A1AT washout from the medium. We also found that the effect of A1AT can be overturned by addition 1 μM trypsin to the bathing solution. Considering the rate at which INa is inhibited in the presence of A1AT (τ = 13 minutes), and the restitution of the current by trypsin, it is consistent with the notion that A1AT acts by inhibiting serine proteases needed for the activation of silent ENaC channels at the cell surface.
It is interesting to note that, although aprotinin has been shown to decrease amiloride-sensitive short-circuit currents across confluent monolayers of HBE cells (10), mouse alveolar epithelial cells (1), and A6 cells (27), it had no effect on baseline rat and mouse Na+-dependent AFC (although it prevented the terbutaline increase of AFC ). In contrast, A1AT decreased baseline AFC in both rats (47) and C57BL/6 mice (present data). One difference between the studies of Planes and colleagues (1) and ours is the baseline values of AFC in C57BL/6 mice. Planes and colleagues reported values of 17% of instilled volume per 30 minutes, whereas our AFC values (consistent with our previous work [20, 37, 48]) are closer to 30% of instilled volume per 30 minutes. In our experiments, A1AT decreased AFC to 12%, which is similar to the baseline value of Planes and colleagues (1).
The structural properties that allow A1AT to inhibit proteases are susceptible to post-translational modifications by oxidation, nitration, and polymerization (49, 50). Neutrophils and macrophages, which are capable of secreting high levels of oxidants at sites of inflammation, have been shown to oxidize two of the nine methionines in A1AT (358 and 351), leading to loss of antielastase activity (51). Similarly, exposure to ONOO− was shown to result in loss of A1AT activity (29). Our data clearly show that ONOO−-treated A1AT does not decrease ENaC function, and suggest that its effects on ENaC are mediated via inactivation of a resident protease present on the cell membrane of oocytes, H441 cells, and on the apical membrane of lung epithelial cells in vivo. Increased levels of reactive oxygen nitrogen intermediates are present in the alveolar spaces of patients with a number of inflammatory diseases (52, 53), which may result in post-translational modifications (such as oxidation and nitration) of important lung proteins (52). Decreased levels of A1AT have also been observed in the lungs of animals exposed to cigarette smoke (54). Decreased levels of native A1AT may be beneficial in some diseases, such as acute lung injury, in which excess amounts of fluid may be present in the alveolar space, which can be cleared via osmotic forces generated by active, vectorial reabsorption of Na+ ions. On the other hand, decreased levels of A1AT in CF may contribute to the pathogenesis of the disease, providing the basis for A1AT administration.
In addition, at the cellular level, A1AT, independently of its protease inhibitory activity, was found to inhibit endotoxin (LPS)-stimulated TNF-α expression and release, and enhance IL-10 expression in human monocytes and endothelial cells by signaling though the antiinflammatory cAMP pathway, which is shared by a number of antiinflammatory drugs currently under development (55). Presently, A1AT is administered to patients with CF to restore protease–antiprotease balance and to reduce lung inflammation (56).
Taken together, the activation of channels by proteases at the apical membrane may be blocked by specific inhibitors, producing a time-dependent reduction in the number of active channels that results from internalization and degradation of already activated channels (27). Our data confirm previous findings on the importance of protease–antiprotease balance in vitro, and offers new insights as to the regulation of ENaC by A1AT, the most abundant circulating antiprotease in vivo. They also highlight a potential mechanism by which A1AT administration may prove beneficial in clinical situations in which increased ENaC activity may contribute to lung pathology.
The authors are grateful to Dr. Peter Snyder, University of Iowa School of Medicine, for the generous gift of Liddle epithelial Na+ channel cRNAs, and to Ms. Teri Potter for editorial assistance in the preparation of this manuscript.
This work was supported by NIH grants HL031197, HL051173, and 5U01ES015676 (S.M.), and grants from the Swedish Medical Research Council, Crafoords Foundation, MAS Foundation, and Grifol's Institute (S.J.).
Originally Published in Press as DOI: 10.1165/rcmb.2008-0384OC on January 8, 2009
Conflict of Interest Statement: S.M. and S.J. received a grant from Talecris Biotherapeutics, Inc., which manufactures Prolastin. S.M. was paid $2,000 to attend the Sepracor Scientific Research Forum at the Windsor Court Hotel (New Orleans, LA), September 18–20, 2008 (travel costs were also paid for by Sepracor), received a grant from Sepracor, Inc. in the amount of $120,127(direct costs) for March 1, 2007 to February 28, 2008 to study “In Vivo Studies on Effiicacy of Aerosolized Alpha1-Antytrypsin,” has a U.S. Provisional Patent Application (no. 61/052,414, “DPPC Formulates and Methods for Using” [filed May 12, 2008; inventors, E.I. Franses, S.H. Kim, Y. Park, and S.M.), and U.S. Provisional Patent Application (no. 60/573,558: “Methods for Using Pyrimidine Synthesis Inhibitors to Increase Airway Epithelial Cell Fluid Uptake” [filed May 21, 2004; inventors: I.C. Davis, W. Sullender, and S.M.), converted to International Patent Cooperation Treaty application (no. PCT/UC2005/017939); in May 2005, D.S. and S.J. received a grant from Talecris for 2006–2007 for $150,000. None of the other authors has a financial relationship with a commercial entity that has an interest in the subject of this manuscript.