The accelerated loss of lung epithelium through activation of extrinsic apoptosis is believed to play a causative role in lung pathogenesis. Previous investigations have shown that zinc is required to sustain lung epithelial cell viability under stress conditions and that depletion of intracellular zinc predisposes cells to apoptosis. In this investigation, we determined whether intracellular zinc deficiency enhanced the susceptibility of primary, differentiated cultures of human lung epithelium to death receptor-mediated apoptosis, leading to barrier dysfunction. Cultures obtained from multiple donors were exposed to stimuli that provoke death receptor-mediated apoptosis and depleted of intracellular zinc with a zinc-specific chelating agent. Transepithelial resistance, paracellular transport, caspase-8 and caspase-3 activity, and apoptosis were measured. Activation of extrinsic apoptosis or zinc chelation alone resulted in a nominal increase in caspase function and apoptosis without major evidence of barrier disruption. Activation of extrinsic apoptosis in addition to zinc depletion resulted in an abrupt decrease in transepithelial resistance, a substantial increase in apoptosis, and an increased paracellular leak. Cultures were rescued by supplementation with zinc sulfate. Further analysis revealed that exogenous zinc facilitates cell survival through activation of the phosphatidylinositol 3-kinase/Akt signaling pathway. We conclude that intracellular zinc is a vital factor in lung epithelium that protects cells from death receptor-mediated apoptosis and barrier dysfunction.
- programmed cell death
- airway epithelium
- barrier dysfunction
destruction of the epithelial barrier at the onset of inflammatory lung disease is a defining event in pathogenesis (9). Extensive epithelial cell loss promotes pathogenic remodeling and alteration of normal tissue architecture, leading to deficits in pulmonary function. Although much is known about elaboration of the inflammatory response in the lung, less is understood about the initial events that trigger epithelial cell loss as well as the environmental or inherited factors that predispose the epithelium to programmed cell death.
The normal airway epithelium is differentiated and stratified and consists of a columnar layer of ciliated and secretory cells supported by basal cells. The polarized structure constitutes a physical barrier that protects the lung from the external environment. Maintenance of this barrier is critical to host defense and normal tissue function. Epithelial damage and desquamation are characteristic of lung disease associated with inflammation and remodeling, including diseases such as asthma and acute respiratory distress syndrome. Recent studies have revealed that programmed cell death of the epithelium is a characteristic feature of patients with established disease (3, 12, 13). This observation has generated interest in further elucidation of the mechanism(s) responsible for lung epithelial apoptosis that creates a foundation for inflammation and pathogenic remodeling. In our initial studies with differentiated, polarized cultures of human airway epithelia, we observed that epithelial barrier function remains intact in conditions of inflammatory stress and that the lung epithelium is refractory to mediators that initiate extrinsic apoptosis. On this basis, we questioned whether alteration of other commonly occurring biological factors associated with cellular homeostasis could contribute to epithelial cell demise in response to apoptotic stimuli.
We hypothesized that intracellular zinc depletion would augment the epithelial response to activators of the extrinsic apoptosis pathway, resulting in cell death and barrier disruption. We based our hypothesis on previous investigations that identified a causal link between intracellular zinc concentration and lung epithelial apoptosis (15, 16). In this report, we provide new evidence demonstrating that intracellular zinc is an important cofactor in the lung epithelial response to activators of extrinsic apoptosis. In contrast to our initial findings, we observed that lung epithelium does not tolerate an inflammatory environment in the absence of zinc. In pursuit of a rationale to explain our results, we provide preliminary evidence that zinc facilitates cell survival through modulation of the phosphatidylinositol 3-kinase (PI3K) survival pathway. We believe that further investigation in this area will generate mechanistic insight into the critical role that dietary zinc plays in maintaining the lung epithelium and, thereby, preserving barrier integrity and will provide insight at the molecular level into innovative therapeutic strategies to treat lung disease.
MATERIALS AND METHODS
Primary cell culture.
Primary human lung epithelial cells were isolated after enzymatic dissociation from trachea, bronchi, and bronchioles of adult donor lungs, seeded onto collagen-coated, semipermeable membranes (0.6 cm2; Millicell-HA, Millipore, Bedford, MA), and grown at an air-liquid interface as previously described (10). Results were derived from six different donors obtained during the course of the investigation. Typically, within 96 h after seeding, the airway cells form a confluent culture with electrically tight junctions and transepithelial resistance >800 Ω. Between days 4 and 14, the epithelial cells differentiate into a predominantly ciliated phenotype. In all experiments, primary cultures were ≥2 wk past initial seeding. Human lung epithelial cells were maintained in 1:1 DMEM-Ham's F-12 medium supplemented with 2% Ultroser G (BioSepra, Villeneuve, La Garenne, France) and antibiotics unless otherwise stated. Human lungs were collected with approval from The Ohio State University Institutional Review Board. For all experiments, unless indicated, the cells were cultured under serum-free conditions for 48 h; then IFN-γ (250 U/ml; catalog no. PHC 4031, BioSource, Camarillo, CA), TNF-α (100 ng/ml; Knoll Pharmaceuticals, Whippany, NJ), and anti-human CD95 (200 ng/ml, Fas Ab; catalog no. MAb 142, R & D Systems, Minneapolis, MN) were added for an additional 48 h. To achieve zinc deprivation, the cultures were grown in non-zinc-supplemented DMEM for 48 h before cytokine addition. To deplete intracellular zinc stores, the zinc chelator N,N,N′,N′-tetrakis(2-pyridylmethyl)ethylenediamine (TPEN, 25 μM) (20) was added 4 h before the end of the 48-h incubation period with cytokines. All treatment conditions were done in triplicate, and each data set is representative of at least three separate experiments unless otherwise stated.
Generation of cell extracts.
After the treatments described above, adherent cells were trypsinized and pooled with nonadherent cells present in the original medium. The cells were pelleted at 300 g for 10 min at room temperature, the supernatants were discarded, and the cell pellet was resuspended in 200 μl of 1× KPM complete buffer (2× KPM = 100 mM PIPES, pH 7, 100 mM KCl, 20 mM EGTA, and 3.84 mM MgCl2), 1 mM DTT, 0.1 mM PMSF, 10 mg/ml cytochalasin B, 2 mg/ml chymostatin, 2 mg/ml leupeptin, 2 mg/ml antipain, and 2 mg/ml pepstatin A. Resuspended cells were transferred to 0.5-ml Eppendorf tubes and pelleted again (3,000 rpm for 10 min). The cell pellets were resuspended in 15 μl of 1× KPM complete buffer, frozen in liquid nitrogen, placed in a room-temperature water bath until cells had thawed, and then vortexed. The freeze-thaw procedure was performed a total of four times. Samples were then centrifuged at 14,000 rpm for 20 min at 4°C. The resulting supernatant or cell extract was removed to a fresh 0.5-ml Eppendorf tube, and an aliquot was analyzed for protein concentration by the Bradford method (Bio-Rad, Hercules, CA), frozen in liquid nitrogen, and stored at −70°C.
Measurement of caspase activity.
Enzymatic caspase activity was measured with 7-amino-4-(trifluoromethyl)coumarin (AFC). For all AFC preparations, 3 × 106 cells were collected by centrifugation, washed with KPM buffer, and lysed by four cycles of freeze-thawing as previously described (6). The presence of active caspases was determined by AFC assay using specific fluorosubstrates. Lysates were incubated with Cyto buffer (10% glycerol, 50 mM PIPES, pH 7.0, and 1 mM EDTA) containing 1 mM DTT and 20 μM Asp-Glu-Val-Asp (DEVD)-AFC or Ile-Glu(OMe)-Thr-Asp(OMe)-AFC (Enzyme Systems Products). The release of free AFC was determined using a fluorometer (Cytofluor 4000, Perseptive, Framingham, MA; 400-nm excitation and 505-nm emission).
Analysis of apoptosis.
Cells were detached using nonenzymatic cell dissociation solution (Sigma, St. Louis, MO), pooled with cells already suspended during culture, and plated onto Silane-treated slides by cytospin. The cells were rinsed twice with 1× PBS, and fixed in ice-cold pure methanol for 30 min at −20°C. After they were washed twice with washing buffer (1× PBS and 0.1% Tween 20), the wells were blocked with blocking buffer (1× PBS, 1% BSA, and 0.1% Tween 20) for 10 min. The cells were then incubated with M30 (CytoDEATH) antibody (Boehringer Mannheim, Indianapolis, IN), a monoclonal antibody that specifically detects caspase-cleaved human cytokeratin-18 (CK-18), diluted 1:10 in blocking buffer for 1 h at room temperature (4). The cells were washed twice with washing buffer and then incubated with 10 mg/ml anti-mouse IgG labeled with fluorescein (Boehringer Mannheim) for 30 min at room temperature in the dark. Cells were again washed twice with washing buffer before incubation with 0.5 mg/ml 4′,6-diamidine-2′-phenylindole dihyrochloride (DAPI; Roche Molecular Biochemicals, Indianapolis, IN) for 5 min at room temperature in the dark. Cells were washed three times with washing buffer and allowed to air dry slightly. Anti-fade (Molecular Probes, Eugene, OR) was added to each slide before coverslips were applied. Cells were considered to be apoptotic if caspase-cleaved CK-18 appeared in the cytoplasm in conjunction with nuclear fragmentation as detected by DAPI staining. Specificity was confirmed by comparison with an antibody that recognizes native CK-18 in all cells as well as comparison with an isotype control antibody, MOPC21 (Sigma). Apoptotic (M30-positive) and total (DAPI-stained nuclei) cells were enumerated by a blinded observer who randomly selected six fields of view per treatment condition. Data are presented as the average percentage of apoptotic cells divided by the total number of cells per viewing area. Results are compared with actinomycin D-treated cells as a positive control where indicated. Lactate dehydrogenase (LDH) release was used as an additional measure of cell death in compliance with the manufacturer's instructions (Roche Applied Sciences, Indianapolis, IN). On the basis of our experience, identical results are obtained with the TdT-mediated dUTP nick end labeling assay, thereby confirming the validity of our assay (6).
Assessment of paracellular transport.
Transepithelial resistance, referred to as TEER or Rt, is monitored using a portable ohmmeter (Millicell-ERS, Millipore). For measurement of Rt, 300 μl of medium were placed on the apical surface and removed after measurement. We also assessed barrier integrity by measuring the paracellular transport of Lucifer yellow as we previously reported (2). In each set of experiments, Lucifer yellow was added to the donor (apical) chamber at a final concentration of 50 μM. At specified time points, 100-μl aliquots of fluid were taken in triplicate from the acceptor (basolateral) chamber and analyzed in a fluorescent plate reader at excitation and emission wavelengths of 425 and 535 nm, respectively. The threshold value to establish tight barriers was a Lucifer yellow flux <0.25%/h. All experimental conditions were compared with a positive treatment control in which inserts were treated for 20 min with medium containing 10 μM EGTA, which transiently opens all junctions between neighboring cells. TEER was monitored in EGTA-treated cells until it fell below 50 Ω, indicating loss of barrier function, which typically occurred within 20 min after the addition of EGTA. In each experiment, EGTA-induced Lucifer yellow flux was set at 100%, and samples were compared with this standard.
Cell surface biotinylation and immunolocalization.
Cell surface biotinylation was carried out as previously described (21). Briefly, the apical surface of each donor culture was exposed to sulfosuccinimidyl-6-(biotinamido)hexanoate (sulfo-NHS-LC biotin), and the cells were lysed. Jurkat cells were used as a positive control and treated identically. Cells on the inserts were dislodged by incubation at 37°C for 30 min with Cellstripper (Mediatech-Cellgro, Herndon, VA), collected using a cell scraper, and washed three to four times with Dulbecco's phosphate buffer (pH 7.4). Biotinylated Fas or TNF receptor type 1 (TNFR1) protein was immunoprecipitated with anti-Fas (anti-human Fas; catalog no. AHS 9552, BioSource, Camarillo, CA) or anti-TNFR1 antibody (anti-TNFR1, catlog no. 654216, Calbiochem, San Diego, CA) and purified before resolution on a 10% Tris·HCl gel (Bio-Rad). An equal amount of immunoprecipitant was loaded for each sample. The resolved proteins were then transferred to a polyvinylidene difluoride membrane and detected with horseradish peroxidase-conjugated streptavidin (Amersham Biosciences, Piscataway, NJ). The same antibodies were used for immunohistochemistry studies to demonstrate cellular staining patterns. Briefly, the cells were nonenzymatically disadhered, plated onto glass slides by Cytospin preparation, and immunostained. A goat anti-rabbit FITC-labeled secondary antibody was used for fluorescent detection (Sigma Immunochemicals, St. Louis, MO).
Detection of PI3K activity and Akt phosphorylation.
After exposure of primary cultures to 100 μM zinc sulfate for 4 h, PI3K was immunoprecipitated from whole cell lysates (catalog no. 06-195, Upstate Biotechnology) and subjected to PI3K-mediated conversion of phosphatidylinositol 4,5-diphosphate to phosphatidylinositol 3,4,5-trisphosphate (PIP3) in a standard kinase reaction based on the manufacturer's recommendations (Echelon Biosciences, Salt Lake City, UT). Samples were also pretreated with the PI3K inhibitor LY-249002 (50 μM) or the receptor tyrosine kinase inhibitor genistein (10 μM) for 1 h before the addition of zinc sulfate. The product was then measured with a competitive ELISA that determines the amount of PIP3 produced. In tandem, we also analyzed whole cell lysates for phosphorylated and total Akt by Western blot.
Values are means ± SE. Paired t-tests were used for single comparisons (Excel, Microsoft, Redmond, WA). For comparisons that involved multiple variables and observations, two- and three-way ANOVA (JMP, SAS Institute, Cary, NC) was used. After statistical significance was established by ANOVA, individual comparisons were made using Tukey's multiple comparison test. Statistical significance was defined as P < 0.05.
Lung epithelium is resistant to extrinsic apoptosis.
Initially, we exposed fully differentiated cultures of primary human upper airway cells to a combination of TNF-α, a Fas cross-linking antibody (Fas Ab), and IFN-γ. Serum-free conditions were utilized throughout all experiments to avoid interference with growth factor-induced simulation of primary cells. Serial measurements of transepithelial electrical resistance (TEER) were recorded immediately before and up to 4 days after reagents were added to the basolateral chamber. Culture inserts grown in serum-free medium maintained high TEER, comparable to serum-supplemented conditions. When TNF-α + Fas Ab + IFN-γ (ITF) was added to the basolateral chamber, a modest reduction in TEER, similar to that observed in serum-free, untreated control chambers up to 4 days after the addition of ITF, was observed (Fig. 1A). Similar results were obtained with different concentrations of the three reagents (data not shown). Despite the reduction in TEER, we observed that the apical surface remained dry, suggesting that barrier integrity was intact. In each experiment, we concomitantly measured apoptosis and a general index of cell viability, i.e., LDH release. Exposure to ITF leads to a small, but significant, increase in apoptosis, as measured by the M30 (CytoDEATH) assay, as well as an increase in LDH release compared with untreated cultures (Fig. 1B).
To determine whether the refractory response to extrinsic apoptosis was due to a lack of expression of the receptors that initiate extrinsic apoptosis, we conducted surface biotinylation-labeling studies on the apical and basolateral surface of polarized, differentiated cultures under baseline conditions. We found expression of the Fas receptor and TNFR1 predominantly at the basolateral surface of fully differentiated upper airway epithelia at their respective molecular masses in cell membrane preparations (Fig. 2A). We also observed expression of the Fas receptor and TNR1 in cytospin preparations (Fig. 2B). Cytospin preparations clearly demonstrate Fas on the cell membrane, whereas TNFR1 was expressed diffusely throughout the cytoplasm as well as on the cell membrane.
Increased apoptosis leads to barrier disruption.
To verify that an increase in apoptosis would lead to larger reductions in TEER and culminate in loss of barrier function, we compared ITF-treated cells with cells treated with actinomycin D, a conventional apoptosis-inducing agent. Actinomycin D treatment resulted in a significant increase in the number of apoptotic cells (Fig. 3A), which correlated with a similar increase in caspase-3 activity as measured by enzymatic function in cell lysates (Fig. 3B). This occurred in tandem with a larger reduction in TEER than was previously observed with ITF treatment (Fig. 3C). We also observed that actinomycin D treatment led to extravasation of culture fluid from the basolateral chamber to the apical surface. To confirm that fluid extravasation was the result of increased paracellular transit and, hence, barrier compromise, we subjected the treatment groups to a Lucifer yellow flux assay. Lucifer yellow is a small 457.2-molecular-weight dye that cannot readily transit through cells or monolayers with physiologically tight junctions. Lucifer yellow transit, however, does occur when the tight junctions between adjacent cells in a monolayer are disrupted. In agreement with our previous findings, only the actinomycin D-treated cells demonstrated substantial paracellular transit of Lucifer yellow across the monolayer, whereas a much smaller degree of paracellular flux occurred in untreated controls or the ITF treatment group (Fig. 3D).
Zinc depletion enhances death receptor-mediated apoptosis.
On the basis of the observation that airway epithelium is refractory to extrinsic apoptosis, we hypothesized that the additional challenge of zinc deprivation would increase susceptibility of the epithelium to programmed cell death. To achieve this experimentally, we cultured cells in a DMEM-based medium without supplemental zinc for 48 h and, in addition, chelated the intracellular labile pool of zinc with TPEN. Treatment with ITF alone generated a nominal amount of apoptosis, comparable to TPEN treatment alone; however, zinc deprivation in addition to ITF treatment led to a substantial increase in apoptosis slightly greater than that induced by actinomycin D treatment (Fig. 4A). We also measured caspase-8 (Fig. 4B) and caspase-3 (Fig. 4C) activity, considered the primary initiator and terminator caspases in extrinisic apoptosis, respectively, in ITF-treated cells compared with cultures deprived of zinc and exposed to ITF. In agreement with our findings, the magnitude of caspase activation was substantially higher in the zinc-deprived, ITF-treated group. We also observed that the decline in TEER was substantially greater, that fluid extravasation occurred, and that Lucifer yellow flux was highest in zinc-deprived, ITF-treated cells (Fig. 4D). The level of apoptosis and barrier dysfunction rivaled that induced by actinomycin D treatment. We then determined whether zinc supplementation could prevent apoptosis and barrier disruption and whether caspase-3 was involved in this process. We found that zinc supplementation protected zinc-depleted, ITF-treated cells from apoptosis and barrier disruption (Fig. 5A). We also observed that blockade of caspase-3 partially inhibited apoptosis but did not prevent barrier rupture (Fig. 5B). We believe this to be in part attributed to toxicity associated with prolonged exposure to caspase inhibitors (unpublished observation). The protective effect rendered by zinc appeared to be relatively specific, because supplementation with magnesium chloride or calcium chloride at concentrations an order of magnitude greater than zinc concentration did not prevent apoptosis (Fig. 5C). Exposure to 100 μM zinc under these conditions did not lead to an increase in LDH release or a substantial decrease in TEER, indicating that zinc exposure alone was not toxic to cells (data not shown).
Zinc induces the PI3K signaling pathway.
Having shown that zinc inhibits extrinsic apoptosis, we wanted to determine whether zinc protects the epithelium by activating PI3K/Akt signal transduction, a pathway that promotes cell survival. First, we subjected zinc-starved cells to zinc sulfate in culture under serum-free conditions and analyzed whole cell lysates for phosphorylated Akt. Untreated cells exhibited evidence of phosphorylated Akt that was substantially increased above baseline within 2 h after cultures were exposed to 100 μM zinc sulfate (Fig. 6A). We also observed that phosphorylated Akt induction was inhibited by an equimolar concentration of TPEN (Fig. 6B). Because PI3K leads to Akt phosphorylation, we analyzed zinc-treated cell extracts for evidence of PI3K activity. To accomplish this, we immunoprecipitated PI3K from zinc-starved cultures 4 h after zinc supplementation and then subjected the immunoprecipitants to a kinase assay followed by analysis with a quantitative ELISA specific for the phosphoinositide product PIP3. We observed that zinc induced PI3K activity (Fig. 6C) and that this activity was inhibited in zinc-treated cells that were also exposed to the PI3K inhibitor LY-249002 (Fig. 6D). Interestingly, we also observed inhibition of zinc-induced phosphorylated Akt by the receptor tyrosine kinase inhibitor genistein, indicating that zinc-induced PI3K activation may act through upstream mediators.
To determine whether zinc-mediated PI3K activity is responsible for zinc-induced cell survival, we exposed cells to a combination of ITF and TPEN and rescued cells with zinc supplementation (Fig. 7). As an additional intervention, we treated cells with LY-249002 immediately before the addition of zinc. PI3K inhibition reversed the protection provided by zinc supplementation in primary differentiated airway epithelial cells, and inhibition was not complete. We believe that this observation suggests that zinc-mediated epithelial cell protection involves PI3K and that other protective mediators are involved. LY-249002 alone did not cause cell death, as measured by M30 staining, nor did it lead to a precipitous decrease in resistance measurements (data not shown).
In agreement with our previous work, we observed that differentiated cultures of primary human upper airway epithelia are susceptible to apoptosis when exposed to a combination of IFN-γ and ligands for TNFR1 and Fas/CD95, receptors that initiate extrinsic apoptosis (6). As expected, these factors induced extrinsic apoptosis mediated through activation of caspase-8 and caspase-3, resulting in cellular apoptosis. However, to our surprise, the magnitude of cell death in fully differentiated human cultures in response to these inflammatory mediators was quite low after extended exposure (as long as 96 h). Although apoptosis was accounted for in the adherent and suspended cell populations after exposure, the percentage of apoptotic cells within the entire culture was typically <10%. Furthermore, despite evidence of apoptosis, the barrier function of the differentiated monolayers remained largely intact. The poor response to these stimuli could not be attributed to a lack of expression of cognate death receptors, because TNFR1 and Fas were abundant and present at the membrane surface. This finding suggested that other factors must be involved in sequestering the cellular response to apoptotic stimuli. On the basis of these observations, we explored the possibility that environmental factors are involved in determining the susceptibility of the lung epithelium to extrinsic apoptosis. To test this, we introduced an additional challenge, i.e., zinc starvation, to our in vitro model.
Zinc, an essential dietary element critical for growth and tissue repair, is known to play an important role as a cytoprotective agent for many cell types, including the lung epithelium, against different noxious stimuli (17, 20). The intracellular pool of exchangeable zinc is naturally abundant in the cytoplasm of upper airway epithelia compared with other cell types and is critical in protection against oxidant-induced apoptosis and allergen challenge. In a murine model of ovalbumin-induced lung allergic hypersensitivity, zinc deprivation enhanced airway inflammation and epithelial damage (16). Similarly, brief exposure to aerosolized zinc sulfate protected guinea pigs against allergic bronchoconstriction and prevented most pulmonary complications associated with this model (5). In our investigation, we observed that zinc deprivation led to a substantial increase in lung epithelial susceptibility to extrinsic apoptosis and contributed to a loss of barrier function. This effect was primarily observed in cultures exposed to inflammatory stimuli, and not in cultures subjected only to zinc deprivation. Furthermore, zinc supplementation inhibited cell death induced by ITF + TPEN, thereby preserving barrier function. This finding raises the possibility that dietary zinc intake or lack thereof may be a contributing factor in determining host susceptibility to inflammation and parenchymal tissue destruction. Similar to previously published reports, we observed that the cytoprotective effect of supplemental zinc on differentiated airway epithelial cells was specific and not reproducible with other commonly occurring divalent cations. Taken together, these observations would indicate that zinc is an important intracellular regulatory factor capable of facilitating cytoprotection against a broad range of inflammatory stimuli, including oxidants, cytokines, and environmental allergens.
The average zinc concentration in a eukaryotic cell has been estimated to be 10−13–10−9 M (7), and strict regulation of cellular zinc content through zinc transporter proteins is required to maintain homeostasis. Approximately 90% is defined as the “stable pool,” which is constant and poorly exchangeable; the remaining 10% is identified as the “labile” pool, which is rapidly depleted in zinc deficiency and readily chelatable under experimental conditions such as ours. The cytoplasmic zinc pool in the lung epithelium is involved in the regulation of many organ-specific, zinc-dependent processes, including signal transduction and apoptosis (15, 16, 19). On the basis of our findings, we believe that the intracellular pool of accessible zinc plays an important role as a cytoprotective agent that facilitates lung epithelial cell survival, particularly during intervals of inflammatory stress. The mechanism by which zinc mediates cytoprotection is likely multifactorial and involves the activation of intracellular signaling pathways associated with cell survival, including the PI3K/Akt signaling axis (11, 19). In support of this theory, supplemental zinc has the ability to stimulate multiple signaling pathways, including Src-dependent growth factor activation of ERK1/2, PI3K-mediated activation of Akt, and the S6 kinase, and to inhibit phosphatases that inactivate these pathways (18). It may also have a role in caspase inhibition. However, a substantial gap exists in our understanding of how zinc directly interacts with these pathways. It is not clear whether the activation of different signaling pathways shares a common induction event coupled to zinc entry or operates through entirely different mechanisms. A zinc-sensing receptor that links extracellular zinc to intracellular signaling events leading to PI3K-dependent ERK1/2 activation has been reported (1, 8, 14); however, there is no direct physical evidence of this receptor. Our investigation supports the notion that a proximal receptor-mediated signaling event is responsible for PI3K activation, in that genistein, a broad-spectrum receptor tyrosine kinase inhibitor, inhibited PI3K function and Akt phosphorylation. We also observed a similar suppression of zinc-induced ERK1/2 activation (unpublished observation).
Our investigation provides new evidence for the relation between zinc, human airway epithelial cells, and regulation of the epithelial response to inflammatory stimuli involved in lung pathogenesis. We observed that the lung epithelium is inherently resistant to extrinsic apoptosis and that barrier function is maintained under a simulated condition of inflammatory stress. In sharp contrast, we found that zinc deprivation substantially increases susceptibility to extrinsic apoptosis and facilitates barrier disruption. This finding supports the concept that host predisposition to acquired lung disease is determined by inflammation in addition to environmental cofactors that regulate the cellular response to inflammatory stress. Our findings suggest that dietary zinc intake, as well as genetic factors that regulate intracellular zinc stores, may be critical components of lung epithelial cell homeostasis and deserve further study. We predict that subacute zinc deficiency may place the host at a significant disadvantage during periods of inflammatory stress that promote lung epithelial cell apoptosis, barrier dysfunction, and pathogenic tissue remodeling. We also propose that zinc-induced protection against extrinsic apoptosis is achieved, at least in part, through activation of the PI3K signaling axis, although the mechanism responsible for kinase activation remains unknown. The supportive contribution of zinc to innate immune defense in the lung has great appeal in terms of helping identify at-risk populations and devising alternative therapeutic strategies through dietary supplementation or topical delivery to prevent barrier compromise and lung disease.
This work was supported by National Heart, Lung, and Blood Institute Grant HL-56336 (D. L. Knoell) and the American College of Clinical Pharmacy (D. L. Knoell).
We thank Dr. Mark D. Wewers for critical insight and manuscript preparation.
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.
- Copyright © 2006 the American Physiological Society