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Am J Physiol Lung Cell Mol Physiol 295: L708-L717, 2008. First published August 8, 2008; doi:10.1152/ajplung.00536.2007
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Long-term exposure to LPS enhances the rate of stimulated exocytosis and surfactant secretion in alveolar type II cells and upregulates P2Y2 receptor expression

Ignacio Garcia-Verdugo,1 Andrea Ravasio,2 Elvira Garcia de Paco,1 Monique Synguelakis,1 Nina Ivanova,3 Jean Kanellopoulos,1 and Thomas Haller2

1Institut de Biochimie et Biophysique Moléculaire et Cellulaire, University of Paris-Sud, Orsay, France; 2Department of Physiology, Innsbruck Medical University, Innsbruck, Austria; and 3Institute of General Physiology, University of Ulm, Ulm, Germany

Submitted 28 December 2007 ; accepted in final form 3 August 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Bacterial LPS is a potent proinflammatory molecule. In the lungs, LPS induces alterations in surfactant pool sizes and phospholipid (PL) contents, although direct actions of LPS on the alveolar type II cells (AT II) are not yet clear. For this reason, we studied short- and long-term effects of LPS on basal and agonist-stimulated secretory responses of rat AT II by using Ca2+ microfluorimetry, a microtiter plate-based exocytosis assay, by quantitating PL and 3H-labeled choline released into cell supernatants and by using quantitative PCR and Western blot analysis. Long term, but not short term, exposures to LPS led to prolonged ATP-induced Ca2+ signals and an increased rate in vesicle fusions with an augmented release of surfactant PL. Most notably, the stimulatory effect of LPS was ATP-dependent and may be mediated by the upregulation of the purinergic receptor subtype P2Y2. Western blot analysis confirmed higher levels of P2Y2, and suramin, a P2Y receptor antagonist, was more effective in LPS-treated cells. From these observations, we conclude that LPS, probably via Toll-like receptor-4, induces a time-dependent increase in P2Y2 receptors, which, by yet unknown mechanisms, leads to prolonged agonist-induced Ca2+ responses that trigger a higher activity in vesicle fusion and secretion. We further conclude that chronic exposure to endotoxin sensitizes AT II to increase the extracellular surfactant pool, which aids in the pulmonary host defense mechanisms.

endotoxin; lipopolysaccharide; lung; P2Y2

PULMONARY SURFACTANT PREVENTS alveolar collapse and aids in the clearance of inhaled pollutants and pathogens. Its components (lipids and proteins) are synthesized and secreted by alveolar type II cells (AT II) via regulated fusion of lamellar bodies (LB) with the plasma membrane (PM). Many agonists have been described to activate LB-PM fusions and to release phospholipids (PL) in isolated AT II (1). Among these, nucleotides such as ATP are likely to be the most physiologically relevant ones (29). ATP exerts its effects through interactions with purinergic receptors, in particular with the G protein-coupled receptor P2Y2 (37), which results in Ca2+ mobilization and activation of PKC. On addition of ATP to cultured AT II, the first LB-PM fusion events initiate after the cytoplasmic calcium concentration ([Ca2+]c) increases above a low threshold level (23). Calcium is considered as the major second messenger to trigger LB-PM fusions and, importantly, to expand the exocytotic fusion pore to facilitate extrusion of the surfactant lipoprotein complex (15).

Bacterial LPS, or endotoxin, is a potent proinflammatory molecule able to induce, in extreme cases, endotoxic shock (25). After inhalation of endotoxin, an increase of inflammatory mediators (e.g., TNF-{alpha}, IL-1β) in bronchoalveolar lavage (BAL) has been reported (40), which might reflect the inflammatory status of the alveolar epithelium after contact with it. In addition, exposure to endotoxin induces alterations of surfactant pool sizes and PL contents in BAL. Fehrenbach et al. (17) described the formation of giant LB and a reduction in PL content in BAL in isolated rat lungs perfused with LPS. Similarly, Viviano et al. (42) reported an increased pool of intracellular surfactant PL but a decrease in extracellular surfactant 72 h after intratracheal LPS administration. Taken together, these data indicate that LPS alters surfactant metabolism and correspond to the surfactant deficiencies associated with the acute respiratory distress syndrome secondary to infections (21).

Mechanisms underlying the alterations of surfactant constituents in BAL after LPS challenge could be explained by effects of proinflammatory mediators secreted by macrophages and lymphocytes on AT II (3, 7) or, conversely, by LPS interacting with AT II directly (2, 11). Because AT II express functional LPS receptors like Toll-like receptor-4 (TLR-4) (4), it is reasonable to study those possible direct effects. Indeed, incubation of cultured rat AT II with S-LPS from Escherichia coli led to enhanced PL secretion that was stimulated by unknown mechanism and independent of PKC (2, 36). Benito et al. (8) showed that intracellular calcium is involved in the stimulatory response to LPS, which let these authors suggest that LPS causes a release of calcium from internal stores.

Along with its capability to regulate surfactant secretion, probably by paracrine mechanisms (33), extracellular ATP promotes immunological functions. A cross talk between LPS and ATP has been described (reviewed in Ref. 18): treatment of monocytes with LPS led to increased amounts of pro-IL-1β and pro-IL-18 but only few amounts of the mature cytokines were released. However, on interaction of ATP with the ion channel-forming ATP receptor P2X7, there was a dramatic increase in the release of mature IL-1β and IL-18. This cross talk might be especially relevant under pathological conditions when the alveolar epithelium is continuously exposed to LPS, like in pneumonia or in cystic fibrosis. In the latter, mucus is trapped in the airways, inducing a chain of events resulting in chronic lung infections caused by opportunistic pathogens. Interestingly, in patients with cystic fibrosis or with pneumonia, alterations in surfactant levels and surfactant composition in BAL fluid has been reported (9, 11, 30, 38).

Here, we tested the hypothesis that exposure to LPS modifies surfactant secretion in AT II directly. For this purpose, we utilized a novel approach to measure the rate and extent of LB-PM fusions in cells grown in microplates (44). Furthermore, we applied Ca2+ microfluorimetry to observe changes in [Ca2+]c due to LPS and developed an enzymatic protocol to quantify the amount of secreted PL along with standard methods using 3H-labeled choline. Our data show that long term, but not short term, incubations with LPS enhances the ATP-induced rate in LB-PM fusions in conjunction with prolonged [Ca2+]c responses, which can be explained by increased P2Y2 expression. Consistent with these intracellular effects, LPS enhanced the basal and, in particular, the ATP-induced release of PL. We suggest that chronic exposure to endotoxin sensitizes AT II with respect to paracrine purinergic stimulation, which contributes to enhanced pulmonary host defense mechanisms.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell preparation and fluorescence measurements. All cell preparations were conducted in conformity with the Austrian rules for animal care and testing. A license from the Austrian government (BMFW: A07/3682) has been granted to T. Haller. AT II were isolated from male Sprague-Dawley rats (180–200 g) according to the procedure of Dobbs et al. (16). Cells were plated and cultured in sterile 96-well tissue culture plates for 24 h in DMEM supplemented with 24 mM NaHCO3 and 10% FCS. Cells were maintained at 37°C, 5% CO2, and humidified conditions in an incubator. Number of cells and their homogeneous distribution between single wells was measured by recording the calcein fluorescence after a 30-min incubation with 1 µM calcein acetoxymethyl ester (AM) at 37°C, followed by washout with Ringer solution. These measurements have been conducted after completion of the respective experiments.

Cells were exposed to LPS overnight (approximately 12–14 h) within 1–2 days after isolation. We found that purity of the cell preparation continued to improve over time, and, after 48 h in culture, purity was >95%. The fluorescence microplate assay for exocytosis was performed as described in great detail in a previous publication (44). In short, cells were loaded with 500 nM LysotrackerGreen DND-26 (LTG) for 30 min and then washed with Ringer solution. The wells containing loaded cells were immediately filled with 100 µl of Ringer containing 2 mg/ml Brilliant Black (BB) and inserted into a fluorescence microplate reader to obtain a cell number-dependent LTG signal for time 0, which then was converted to 100%. BB served to block extracellular fluorescence due to loss of LTG from fused vesicles as previously described (44). Test substances, all dissolved in 2 mg/ml BB, were applied outside the instrument, after which a kinetic protocol was started. Data were analyzed as previously described (44). All microplate measurements were performed in triplicate. After completion of the exocytosis assay, cells were washed with culture media, and cell viability was accessed by a resazurin reduction assay (27).

Intracellular Ca2+ measurements. Coverslips with adherent fura 2-AM-loaded cells (1 µM, 15 min at 37°C) 2 days after isolation were mounted on the stage of an inverted microscope (Zeiss 100) equipped for epifluorescence measurements and image analysis (TILL Photonics), and measurements were performed as described elsewhere (23).

Quantitation of released PL. Cells were grown in 24-well plates in DMEM + 10% FCS up to 90% confluence. At day 2 after isolation, cells were washed in Ringer solution, and 250 µl of fresh solution containing secretagogues and/or LPS were added. After incubation at 37°C in a CO2 incubator (2 and 5 h, respectively), supernatants were centrifuged and analyzed for PL content using coupled enzymatic reactions based on a previous method to measure PL in serum (31). To 100 µl of supernatant transferred into a 96-well, we added 100-µl "reaction mix" containing phospholipase D (1 U/ml), choline oxidase (0.2 U/ml), horseradish peroxidase (HRP; 2 U/ml), and Amplex Red (0.1 mM), and kinetic of the reaction was recorded with a plate reader by monitoring resorufin formation ({lambda}excitation = 540 nm, {lambda}emission = 595 nm). End points were taken after 90 min at 37°C. In addition, 200 µl of each supernatant was submitted to hydrophobic extraction according to the method of Bligh and Dyer (10). Chloroformic fractions were dried in tubes, and reaction mix was added directly to the dried lipid film and incubated at RT for 90 min under vigorous shaking. Afterward, solutions were transferred into 96-well plates and measured as described before. {alpha}-Lecithin was used in both approaches as PL standard. Scintigraphic determination of PL was done by including [3H]choline chloride (1 µCi per well in 12-well plates) to the medium (DMEM without choline + 10% FCS) for 24 h with/without LPS, 1 day after isolation. Thereafter, cells were rinsed 3 times with serum-free DMEM, and new medium was added, containing no or 100 µM ATP, respectively. Media were removed after 2 or 5 h, and cells were rinsed with fresh media. The samples were combined, and cells were removed by centrifugation. Attached cells were lysed in cold 0.1% Triton X-100. Hydrophobic extractions were performed from supernatants and lysed cells by the method of Bligh and Dyer (10). The extracts were dried, and radioactivity was measured in a β-counter 1209 Wallac (Pharmacia). Secreted PL are expressed as the amount of radiolabel in the medium relative to the total amount (cells + medium) by 100%. Lactate dehydrogenase activity, measured as previously described (5), revealed that none of the agents tested had a significant effect on cell viability or PM integrity.

Real-time PCR assays. Total RNA (2 µg), extracted from cells using TRIzol reagent (Invitrogen), was reverse-transcribed using SuperScript First-Strand Synthesis System for RT-PCR (Invitrogen). The real-time PCR reactions were performed using LightCycler FastStart DNA MasterPlus SYBR Green I mix (Roche) in a LightCycler 2.0 (Roche) according to manufacturer's instructions. The sequence of rat P2Y2 primer pairs used correspond to forward 5'-TTCTCTAGAGCGTGGACCTC-3' and reverse 5'-GACTGAGGCAGGAAACAGGA-3'. Primer pairs for rat β-actin were described previously (45) and correspond to forward 5'-ATCGCTGACAGGATGCAGAAG-3' and reverse 5'-TCAGGAGGAGCAATGATCTTGA-3'. Primers were chosen to result in amplicons of 174 bp that span intron-exon boundaries to exclude genomic DNA contamination. Agarose gel electrophoresis of the P2Y2 product revealed a single product of the expected size. The cDNA was amplified according to the following steps: 1) 95°C for 10 min; 2) 95°C for 10 s; 3) 62°C for 10 s; and 4) 72°C for 20 s. Steps 2–4 were repeated for an additional 39 cycles. At the end of the last cycle, temperature was increased from 65 to 95°C (0.1°C/s) to produce a melt curve. Each PCR product showed a single peak in the melt curves. Standard curves for real-time PCR protocols with all primer pairs obtained with sequential dilutions of one cDNA sample were found optimal (amplification efficiency ≥95%). Analyses of P2Y2 receptor expression relative to the β-actin endogenous control was performed with LightCycler software in relative quantification mode following manufacturer's instructions and using control cells as calibrator. Mean values ± SE of three independent rat preparations are shown (Fig. 6). PCR reactions for P2X7 and the {gamma}-aminobutyric acid receptor {pi}-subunit were performed according to Ref. 13.


Figure 6
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  		Fig. 6. Expression of purinergic receptor subtype P2Y2 in LPS-treated and control cells. A: P2Y2 receptor RNA relative to the actin was analyzed by real-time PCR using LightCycler software and control cells as calibrator as described in MATERIALS AND METHODS. The difference to control was significant (P < 0.01; n = 3 independent preparations). B: expression of P2Y2 by Western blot. Equivalent amounts of treated (5 µg/ml LPS overnight) and untreated AT II were lysed in sample buffer, electrophoresed, and blotted. P2Y2 was detected in blotted membranes using rabbit anti-P2Y2 antibodies (1:500). Actin was used as loading control. For this purpose, the same membranes were stripped and reprobed with anti-actin antibodies. A representative experiment is shown. C, control. C: 3 independent rat preparations were processed as described above, and P2Y2 and actin reactive bands were evaluated by densitometry. The P2Y2 intensity was referred to the corresponding value of actin. Finally, the relative intensity of P2Y2/actin for LPS-treated cells was referred to P2Y2/actin value for the corresponding control (densitometric unit for control = 1). P < 0.01 (t-test).

 
Western blots. AT II grown to confluence in 12-well plates were lysed in sample buffer (50 µl per well) containing 2% 2-mercaptoethanol and immediately boiled for 10 min. Samples were kept at –20°C for no more than 1 wk. Afterward, samples were boiled again for 10 min and subjected to SDS-PAGE in 10% polyacrylamide gels under reducing conditions, after which the resolved proteins were electro-transferred to nitrocellulose membranes in semidry conditions. The membranes were then sequentially incubated in PBS containing 5% nonfat dry milk overnight, 1:500 dilution of rabbit anti-P2Y2 antibodies (Alomone Labs), 1:500 dilution of rabbit anti-adenosine A2B antibodies (Alomone Labs), or 1:500 dilution of rabbit anti-surfactant protein D (SP-D) antibodies (Santa Cruz) for 2 h, 1:10,000 dilution of HRP-conjugated goat anti-rabbit IgG (Bio-Rad) for 1 h, and ECL Plus chemiluminescence reagent (Amersham Biosciences). The membranes were exposed to X-ray film (Amersham Biosciences). Membranes were stripped and reprobed sequentially with 1:5,000 goat anti-actin antibodies (Santa Cruz), 1:10,000 donkey anti-goat HRP-conjugated antibodies (Santa Cruz), and ECL Plus. The autoradiographs were quantified by scanning densitometry (Molecular Dynamics) and analyzed with ImageQuant software. For each rat preparation (n = 3), the intensity of P2Y2 band was referred to actin value, and therefore LPS-treated cells referred to control cells (densitometric unit for control = 1).

Solutions and reagents. Ringer solution contained, in mM, 140 NaCl, 5 KCl, 1 MgCl2, 2 CaCl2, 5 glucose, and 10 HEPES (pH 7.4 at 25°C). LTG, calcein, fura 2, resazurin, and the Amplex Red Phospholipase D Assay Kit were purchased from Molecular Probes, BB from MP Biomedicals, phospholipase D, DMEM, and suramin from Sigma, FCS from Biochrom, and elastase (for the AT II cell preparation) from Elastin Products. LPS from Salmonella minnesota (Re mutant) was obtained from Sigma. Human SP-A was purified from BAL of patients with alveolar proteinosis as described previously (20).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Effects of LPS on the rate of exocytosis. To investigate if LPS has an effect on exocytosis, i.e., the rate of fusion of intracellular LB with the PM, we used a fluorescence-based microplate assay for adherent AT II as recently described in detail (44). Briefly, a microplate reader collects the emitted light from LTG-loaded cells. This signal correlates with the amount of intracellular LB and declines rapidly when the cells activate these vesicles to release their contents, including the water-soluble LTG, into the extracellular space. An increased rate of vesicle fusions results in a time-dependent deviation of the measured signals between treated cells and untreated controls against an identical background of a linear fall in both groups, which is due to other and unspecific reasons (e.g., constitutive exocytosis, photobleaching; quenching effects; Ref. 44). An example of this assay is given in Fig. 1A, demonstrating the rapid effect of ATP compared with nonstimulated cells and the early termination of the increased vesicle loss within several minutes thereafter. Addition of LPS (5 µg/ml) immediately after the start of the measurement (after time 0) neither increased nor decreased the stimulated rate of exocytosis and also had no effect on unstimulated cells.


Figure 1
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  		Fig. 1. LPS effects on the rate of exocytosis. A: addition of ATP (100 µM) during a microplate measurement (i.e., between 1st and 2nd data points) caused a pronounced acceleration in fluorescence loss compared with untreated controls (CTRL). LPS (50 µg/ml) applied at the same time neither changed the spontaneous nor the agonist-evoked declines in fluorescence intensity. B: preincubation of alveolar type II cells (AT II) with LPS overnight, at 3 different concentrations, markedly enhanced the stimulation by ATP, whereas no effects were observed in nonstimulated controls. C: overnight exposure of cells with LPS (5 µg/ml) enhanced the ATP (100 µM)- and the ATP + PMA (500 nM)-stimulated rates of exocytosis. D: results of all microplate measurements, demonstrating the change in relative fluorescence units (RFU), determined 30 min after start of the measurement, compared with controls. The effects of LPS were all significant (P < 0.10–5 ATP + LPS vs. ATP; P < 0.05 for ATP + PMA + LPS vs. ATP + PMA), except for the nonstimulated controls. Number of experiments (cell preparations) are indicated within the bars, with the number of single measurements (wells) given in parentheses.

 
In cells treated with LPS overnight, however, we observed a significant acceleration in the rate of ATP-stimulated exocytosis (Fig. 1B), which was practically independent of the LPS concentration used. Again, an effect of LPS was not present in the unstimulated control group of cells. In Fig. 1C, cells were stimulated by either ATP or ATP + PMA. Either mode of stimulation is known to induce a strong exocytotic response, however, each by acting via different cellular mechanisms (37). Given in combination, both secretagogues showed an almost additive and maximal effect, whereas the PMA-induced response is more sustained than that of ATP alone and does not terminate even after 30 min of recording (19). Preincubation of cells with LPS overnight enhanced the ATP-induced rate of exocytosis but also the maximum rate obtained with ATP + PMA, and no effect was seen in control cells (Fig. 1B). Results of all microplate measurements, demonstrating the change in relative fluorescence units compared with controls, are shown in Fig. 1D. The effects of LPS were all highly significant (P < 0.10–5 ATP + LPS vs. ATP; P < 0.05 for ATP + PMA + LPS vs. ATP + PMA) except for the nonstimulated cells. SP-A, which inhibits surfactant secretion but not the rate of vesicle fusions (6), had almost no effect (unpublished observation), ruling out the possibility that any corelease of SP-A by AT II interferes with the results of the fusion assay.

Additional experiments of the same kind were performed to investigate the time dependence of incubation of cells with LPS (Fig. 2A), to test for a dose-response curve of LPS (at constant ATP; Fig. 2B), and of ATP (at constant LPS; Fig. 2C), respectively. As shown, an incubation time of cells with LPS of 2 h yielded a statistically significant (P < 0.005) difference compared with nontreated cells, and a maximum effect was obtained between 8 and 24 h (Fig. 2A). The dose-response curve of LPS (Fig. 2B) demonstrates that overnight exposure to AT II had an already significant (P < 0.001) effect at the lowest concentration used (40 ng/ml), and the maximum was obtained at concentrations as low as 80 ng/ml with no tendency to change even up to 25 µg/ml (see Fig. 1B). Both results (Fig. 2, A and B) are in contrast to Romero et al. (36), who found a stimulatory effect up to 200 µg/ml and a maximum secretion between 2–3 h. Nevertheless, in all subsequent experiments, we used LPS at 5 µg/ml, as it is used in most other studies on lung epithelial cells (2), and an incubation time of 12–14 h. The dose-response curve for ATP (Fig. 2C) revealed that LPS exerted a stimulatory effect throughout all ATP concentrations used. However, the stimulatory effect was strictly ATP-dependent and seemed to become more pronounced the higher the ATP concentration. Additional experiments (Fig. 2, right plots) served to rule out the possibility that ATP, LPS, or both in combination have an effect on cell viability, the amount of injured cells or their metabolic state. As can be seen, the rate in reduction of resazurin into resorufin, a sensitive assay for these parameters (27), was essentially the same under all conditions tested.


Figure 2
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  		Fig. 2. Further analysis of LPS effects on ATP-stimulated exocytosis. A: time dependence of LPS preincubation on {Delta}% RFU (i.e., difference of fluorescence signals between time 0 and after 30 min of measurement). Shown are the values for control cells (without LPS preincubation and ATP stimulation), cells stimulated by ATP, and cells preincubated with LPS and stimulated by ATP. B: concentration dependence of LPS preincubation (overnight) on {Delta}% RFU at constant ATP (100 µM). C: dose-response curve for ATP with and without overnight preincubation by LPS (5 µg/ml). As seen, the LPS effect was strictly ATP-dependent. Figures on the right show the viability in % of controls, determined with the resazurin reduction assay, under each condition tested in A–C, respectively. Re-LPS, Re mutant LPS.

 
Effects of LPS on agonist-induced Ca2+ signals. Addition of ATP (100 µM) to adherent AT II caused a transient Ca2+ increase with a rapid decline toward resting levels as shown in Fig. 3. In LPS-preincubated cells (5 µg/ml overnight), this decline was significantly (P < 0.001 at 200 s) prolongated, and resting Ca2+ levels (i.e., before ATP) slightly elevated (P < 0.001). However, the amplitude of the ATP-induced peak in the fura 2 ratio was nearly identical under both conditions. Short-term incubations with LPS (up to 2 h) had no effects (data not shown). According to recent publication (19), an elevated Ca2+ concentration over time is considered to be the major determinant of LB-PM fusions and an important determinant for expanding the exocytotic fusion pores thereby accelerating surfactant release (24).


Figure 3
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  		Fig. 3. LPS effects on intracellular Ca2+ signals evoked by ATP. Addition of ATP (100 µM; between 2nd and 3rd data point = 10–20 s) to AT II caused a transient Ca2+ increase with a rapid decline toward preresting levels. In LPS-preincubated cells (5 µg/ml overnight), this decline was significantly (P < 10–5) prolongated. The inset is an enlarged view to demonstrate the prolongated peak amplitude in LPS-pretreated cells. Data were analyzed from 170 single cells of 4 different lung preparations. Arbitrary units (arb. u) are shown.

 
Effects of LPS on the amount of released PL. The enhancement of ATP-induced Ca2+ signals and the marked increase in the rate of vesicle fusions by LPS certainly suggest an increase in the amount of released surfactant PL. To test this, we applied a sensitive enzymatic approach as described above and for which kinetics using lecithin standards are shown in Fig. 4A. Our data show that overnight exposure to LPS significantly increased PL in the supernatants under all conditions tested, in nonstimulated controls and in cells stimulated 5 h by ATP or PMA, respectively (Fig. 4B). However, in contrast to the results of the fusion assay and to the well-known action of ATP, this secretagogue produced less PL than in controls. Presumably, a compound coreleased by AT II could interfere with one of the enzymatic reactions. We speculated that it might be SP-A because SP-A (but not ATP) exhibited a significant inhibitory effect on resorufin production (Fig. 4C) and because ATP stimulation might release larger amounts of SP-A than does PMA (unpublished observations). Therefore, we repeated these measurements using hydrophobic extracts of cell supernatants to remove interfering, nonlipidic contaminants (Fig. 4D). Under these conditions, the effect seen in Fig. 4C was partially reverted, and we now obtained an augmented release of PL in LPS-preincubated cells, however, in ATP-stimulated cells only (Fig. 4D). Further analysis using background-corrected data revealed this increase also in PMA-treated cells (Table 1).


Figure 4
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  		Fig. 4. Measurement of phospholipid (PL) in cell supernatants by coupled enzymatic reactions leading to the fluorescent product resorufin. A: kinetics of resorufin formation at different concentrations of lecithin. An unspecific increase in background fluorescence (at lecithin = 0) was always present, obviating a clear determination of termination of the reactions. Nevertheless, end points were taken at 90 min, yielding an almost linear concentration-dependency (inset). B: application of this method for the determination of PL in cell supernatants from LPS-pretreated (gray) vs. untreated (black) cells (n = 7) C: effects of ATP (100 µM) and surfactant protein A (SP-A; 20 µg/ml) on resorufin fluorescence measured at various lecithin concentrations. The ATP effect was not significant, whereas SP-A markedly inhibited resorufin formation by ~40%. D: PL determination in hydrophobic extracts of cell supernatants (n = 5). *P < 0.05; **P < 0.01; ***P < 0.005.

 

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  		Table 1. Mean % increase in PL release after LPS treatment

 
To corroborate these findings and circumvent a possible interference with enzymatic PL detection, we applied a standard method using [3H]choline incorporation into AT II and scintigraphic measurement of radiolabel recovered in cell supernatants (14). These measurements (Fig. 5, A and B) revealed an approximately 50–100% increase in secretion by ATP over controls, about an identical but time-dependent increase in only LPS-treated cells, and an almost twofold increase in combined LPS/ATP-treated cells. Furthermore, we repeated these measurements with UTP, a nucleotide known to act on P2Y2 more specifically than ATP (37). The results (Fig. 5C) revealed essentially the same pattern, fostering our hypothesis that LPS is acting on a signaling pathway involving the P2Y2 receptor.


Figure 5
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  		Fig. 5. A–C: measurement of PL in cell supernatants by the [3H]choline incorporation method. Left: secretion expressed as cpmsupernatant x 100/(cpmsupernatant + cpmcells). cpm, counts/min. Right: secretion expressed in % above untreated controls. Paired t-tests (asterisks) were applied between LPS vs. controls and between ATP (UTP) + LPS vs. ATP (UTP). UTP was used at the same concentration as ATP (100 µM), n = 6; *P < 0.05; **P < 0.01; ***P < 0.005. PC, phosphatidylcholine.

 
Effects of LPS on P2Y2 expression. The P2Y2 receptor is certainly one of the most promising candidates in search for a mechanistic explanation of the LPS effects. Indeed, real-time PCR measurements revealed a fourfold increase in the expression of mRNA encoding this purinergic receptor subtype in overnight LPS-treated cells (Fig. 6A). The difference with controls was highly significant in three independent measurements of three different preparations. To also test for a change in receptor protein, we used Western blots and densitometric analyses. The results (Fig. 6, B and C) demonstrated an elevated P2Y2 protein expression after overnight treatment with LPS (P < 0.01).

Effects of suramin on ATP-induced Ca2+ signaling. Suramin, a P2 receptor antagonist, attenuated the ATP-induced Ca2+ signals slightly (Fig. 7). Suramin and ATP were added to the cells in combination at 100 µM each (previous tests revealed that suramin alone has no effect on [Ca2+]c and that preincubation of cells with suramin, for different times at different concentrations, did not show higher inhibitory efficiencies). In control cells, the effect of suramin was a decrease in the peak amplitude of [Ca2+]c (Fig. 7A), whereas no effect was seen with regard to the intensity and duration of the Ca2+ signal (expressed as the integrated fura 2 ratio values over time; Fig. 7B). In contrast, suramin significantly decreased both peak amplitude and duration of the Ca2+ signal in LPS-pretreated cells.


Figure 7
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  		Fig. 7. Depression of ATP-induced Ca2+ signals by the P2Y receptor antagonist suramin (Sur). A: suramin effects on the ATP-induced peak fura 2 ratio in control and LPS-treated cells. *P < 0.05; **P < 0.01. B: suramin effects analyzed with respect to the integrated fura 2 ratio over time (including peak and fura 2 ratio recordings within a 15-min time frame). The suramin effects were all significant except controls vs. suramin in B (n = 3, 9–17 cells each). Shown is the % decrease by treatment with suramin. Note that in these experiments, LPS pretreatment also increased the peak amplitude compared with control cells (A; P < 0.01).

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Alveolar epithelial cells are continuously exposed to microbial challenges. Recognition of invading pathogens is mediated by TLR. Specifically, TLR-4 recognizes LPS and mediates secretion of inflammatory mediators (IL-8 and β-defensin) after challenge of AT II with LPS (4, 28). The surfactant components secreted by AT II, on the other hand, have several pro- and anti-inflammatory properties: surfactant proteins (SP-A and SP-D) are part of our innate immune system. They opsonize pathogens and activate immunocompetent cells (32). In addition, surfactant PL, at high surface pressures in alveoli, permanently feed the mucociliary escalator together with entrapped particles (34). It is reasonable, therefore, to speculate that augmented surfactant secretion is an adaptive response of the alveolar epithelium to a modest LPS challenge, which may be additionally amplified by inflammatory mediators (e.g., TNF-{alpha}, IL-1β) released during aggravation of an infectious state.

Surfactant secretion occurs constitutively and can be further activated by a variety of agonists (1). Nucleotides like ATP are probably the most physiologically relevant agonists, and it is assumed that they primarily act via paracrine mechanisms released during stretch of the alveolar epithelium or by cell damage (33). Both situations may become particularly relevant in pathogen-induced lung hyperinflation, loss of compliance, and associated tissue damage (26), although this issue is still speculative. ATP exerts its effects through interactions with purinergic receptors, in particular with the G protein-coupled receptor P2Y2 (37), which results in Ca2+ mobilization and activation of PKC. In isolated AT II, ATP leads to an initial Ca2+ spike due to Ca2+ release from internal stores, followed by a sustained Ca2+ entry from outside. Numerous studies demonstrate the importance of this second messenger in the regulated secretion of surfactant.

Here, we showed that LPS from S. minnesota increased the ATP-induced fusion of secretory vesicles considerably. This effect was strictly ATP-dependent as it followed in proportion to the ATP concentration used (Fig. 2C). By investigating the Ca2+ signals evoked by ATP, we found a significant prolongation of an elevated [Ca2+]c in LPS-preincubated cells (Fig. 3). This observation is consistent with increased exocytosis and with earlier reports claiming a "Ca2+ dose" above threshold to be essential in some late steps in vesicle fusion. In particular, the amount of fusion pores formed between LB and the PM was found to be strictly correlated to the integrated Ca2+ signal over time (duration x intensity; Refs. 19, 24). In addition, resting Ca2+ levels were also elevated in LPS-treated cells (Fig. 3). However, this did not lead to detectable differences in the spontaneous rate of vesicle fusions (Fig. 1, A, B, and D, and Fig. 2C) but may be the explanation of enhanced lipid transfer into the extracellular space in LPS-treated cells. The reason why this is not reflected by an increased rate in vesicle fusion may be the much shorter time interval (15 min) used in the fusion assay than for measuring PL in cell supernatants (2 and 5 h, respectively). Alternatively, a slightly elevated [Ca2+]c, not sufficient to enhance the constitutive LB-PM fusion, might be essential for the enlargement of the fusion pores of constitutively fused vesicles (Fig. 4, B and D, and Fig. 5). Finally, whether LPS also increased the peak amplitude of [Ca2+]c cannot be stated with certainty. A significant increase (as seen in those experiments of Fig. 7A) was in fact abolished when the data from all experiments were pooled (as in Fig. 3). Accurate peak determinations may be critical as they require much higher sampling rates (e.g., 1 s) than those used in our protocols (10 s). Thus our data only suggest a tendency toward higher peak amplitudes by treatment with LPS.

The above findings are consistent with the observation that intracellular calcium is elevated in sepsis in some tissues (39) and in line with the reports of Benito et al. (8), who showed elevated Ca2+ and an increased phosphatidylcholine secretion due to LPS from E. coli. However, these authors demonstrated that Ca2+ increase occurs within a few minutes after LPS application, whereas our measurements focused on overnight exposures. We have chosen this protocol because immediate effects (<1 min) of LPS could not be detected, neither on intracellular Ca2+ nor on the fusion activity, and by careful examination, a maximum LPS effect was found to peak around 10 h (Fig. 2A). Nevertheless, the measurements of Benito et al. (8) demonstrate a slow decline of [Ca2+]c after LPS exposure, suggesting that Ca2+ might still be elevated even in prolonged exposures. Despite a long-lasting, increased [Ca2+]c due to overnight LPS challenge, our cells did not show signs of reduced viability (Fig. 2), nor did they show obvious signs of abnormalities with respect to number and size of intracellular LB (data not shown), which has been reported earlier (17, 41).

The increase in the rate of vesicle fusions by LPS, and in particular the increase in the amount of PL detected in cell supernatants, are in agreement with other reports (8, 36, 38) but also in contrast to the findings of Fehrenbach et al. (17), who reported a reduction of PL concentration in BAL after LPS challenge, and to Rice et al. (35), who found a depressed AT II cell function in animal models of pneumonia. However, their experimental protocols differed substantially from ours: 1) we used isolated cells, which were exposed to LPS directly, therefore bypassing macrophage-mediated responses and the possible endothelial-epithelial cross talks; 2) we used LPS from a different source; 3) we used longer incubation times; and 4) we primarily investigated the ATP-evoked cell response rather than a constitutive surfactant release. Therefore, we assume that LPS effects may not be comparable between direct and indirect modes of application.

The change in intracellular Ca2+, and in particular the ATP-dependent effect of LPS on vesicle fusions and surfactant secretion, suggested the involvement of a key element in signal transduction, probably the P2Y2 receptor. Real-time PCR data revealed a fourfold increase in the expression of mRNA in LPS-treated cells (Fig. 6A). A change in transcription and insertion of receptors into the PM would fit with the time course of the maximum LPS effect, which was in the range of many hours (Fig. 2A). In line with the PCR data, Western blot analysis revealed a higher level of P2Y2 protein in LPS-treated cells, although a moderate one (Fig. 6, B and C). A decisive proof of increased P2Y2 receptors might have been obtained by blocking specific receptor functions. However, selective inhibitors of P2Y2 are not available: reactive blue, for example, is reported to act as a potent P2 agonist instead of being an antagonist (22). Suramin, a general but unspecific P2 receptor antagonist, had only modest effects on [Ca2+]c (Fig. 7), although it should be more reactive on P2Y (12, 43) than on other purinergic receptor subtypes. Nevertheless, the suramin effect on the Ca2+ peaks seemed to be higher after incubation of cells with LPS (Fig. 7A). Furthermore, LPS-treated cells showed a significantly attenuated Ca2+ dose that was not seen in control cells. These findings tentatively suggest an increased level in functional P2Y2 receptors. This is indirectly supported by the observation that adenosine A2B receptors, a main alternative route of ATP signaling (37), remained unchanged (Supplemental Fig. 2, available in the data supplement online at the AJP-Lung Cellular and Molecular Physiology web site). In light of the weak performance of suramin in our cells, however, these data may only allow the conclusion of a relatively higher contribution of P2Y2-mediated ATP signaling than compared with untreated controls. The data also suggest a major contribution of alternative pathways stimulated by ATP and/or ATP metabolites. This is particularly evident by the fact that LPS did not affect the peak value of ATP-stimulated intracellular calcium, which would be expected in case of increased P2Y2 receptor density, but primarily affected the decay time for the signal. Thus increased expression of P2Y2 may only be an important first evidence to explain changes in signal transduction and Ca2+ homeostasis, and additional mechanisms leading to prolonged Ca2+ signals (e.g., via increased Ca2+ entry or decreased Ca2+ extrusion) still need to be clarified.

P2Y2 receptor upregulation may be specific or may be the result of a general effect of LPS on the preservation of the AT II phenotype in culture. In line with this, we found by real-time PCR an LPS-induced upregulation of the {gamma}-aminobutyric acid receptor {pi}-subunit, an AT II-specific marker (Ref. 13; Supplemental Fig. 1). Although a preserved AT II phenotype would be an alternative hypothesis to explain a higher ATP responsiveness in the present study, several other lines of evidence argue against this. First, expression of the adenosine A2B receptor remained unchanged, as was the case for SP-D (Supplemental Figs. 2 and 3). Similarly, the AT I cell-specific marker P2X7 (13) was not detectable on the transcriptional level, ruling out an increased transdifferentiation of AT II toward AT I in non-LPS-treated cells (Supplemental Fig. 1). Second, LPS effects were obvious after a few hours of incubation (Fig. 2A). Differences between phenotypic preservation between controls and LPS-treated cells are thus very unlikely. To this end, an LPS-induced deceleration of AT II transdifferentiation cannot be entirely ruled out but requires extensive further investigations, including a number of functional and surfactant-related markers.

LPS also seemed to increase the exocytotic response in ATP + PMA-stimulated cells. PMA acts differently than ATP as it bypasses the P2Y2 receptor and directly activates PKC, suggesting that LPS exerts an effect downstream of this receptor. However, several arguments do not support this concept. First, LPS did not significantly increase the amount of PL released from PMA-treated cells (Fig. 4, B and D). Second, the positive effect of LPS in ATP + PMA-treated cells (Fig. 1D) can be explained by its action on the ATP signaling process alone: the percentage decline of ATP + LPS- and ATP-treated cells and that between ATP + PMA + LPS and ATP + PMA was essentially the same. Our assumption is therefore consistent with two earlier reports that LPS acts independently of PKC (8, 36). PMA was used in this study as an additional and independent functional control of the isolated cells and to have an indication of an almost maximum cell response. However, it was not our intention to study effects of this PKC activator further as it is not a physiological agonist like ATP, for which we found, in conclusion, that the endotoxin LPS strongly potentiates its paracrine effects. The signaling pathway involved therein most likely involves an increased expression of functional P2Y2 and a concomitant change in intracellular Ca2+ homeostasis that enables AT II to increase their secretory rate in response to inflammatory processes. However, the transition between direct (by LPS) and indirect effects (e.g., by TNF-{alpha}, IL-1β) of AT II secretion may be a continuous one, and further studies will be necessary to study these effects in combination.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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REFERENCES
 
This work was supported by the Austrian Science Foundation Grant P17501 [GenBank] and the EU Project MRTN-CT-2004-512229 ("Pulmo-Net").


   ACKNOWLEDGMENTS
 
Parts of this work were presented at the "Pulmo-Net" Meeting in Paris, 2006. Technical assistance by I. Öttl and G. Siber is gratefully acknowledged.


   FOOTNOTES
 

Address for reprint requests and other correspondence: T. Haller, Dept. of Physiology and Medical Physics, Division of Physiology, Innsbruck Medical Univ., Fritz-Pregl-Str. 3/1, A-6020 Innsbruck, Austria (e-mail: thomas.haller{at}i-med.ac.at)

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.


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