Apoptosis plays a causative role in acute lung injury in part due to epithelial cell loss. We recently reported that zinc protects the lung epithelium during inflammatory stress whereas depletion of intracellular zinc enhances extrinsic apoptosis. In this investigation, we evaluated the relationship between zinc, caspase-3, and cell-to-cell contact via proteins that form the adherens junction complex. Cell adhesion proteins are directly responsible for formation of the mechanical barrier of the lung epithelium. We hypothesized that exposure to inflammatory cytokines, in conjunction with zinc deprivation, would induce caspase-3, leading to degradation of junction proteins, loss of cell-to-cell contact, and compromised barrier function. Primary human upper airway and type I/II alveolar epithelial cultures were obtained from multiple donors and exposed to inflammatory stimuli that provoke extrinsic apoptosis in addition to depletion of intracellular zinc. We observed that zinc deprivation combined with tumor necrosis factor-α, interferon-γ, and Fas receptor ligation accelerates caspase-3 activation, proteolysis of E-cadherin and β-catenin, and cellular apoptosis, leading to increased paracellular leak across monolayers of both upper airway and alveolar lung epithelial cultures. Zinc supplementation inhibited apoptosis and paracellular leak, whereas caspase inhibition was less effective. We conclude that zinc is a vital factor in the lung epithelium that protects against death receptor-mediated apoptosis and barrier dysfunction. Furthermore, our findings suggest that although caspase-3 inhibition reduces lung epithelial apoptosis it does not prevent mechanical dysfunction. These findings facilitate future studies aimed at developing therapeutic strategies to prevent acute lung injury.
- programmed cell death
- lung epithelium
- barrier dysfunction
- adherens junction
the lung epithelium forms the primary barrier that separates the air space from the fluid-filled vascular and interstitial spaces, thereby providing a high resistance to fluid and solute movement. The pulmonary epithelial barrier not only restricts the passive flow of aqueous fluid and solutes but also contributes to the removal of air space fluid by active transport of solutes across the epithelium from the airway lumen (16). In general, the lung epithelial barrier is much less permeable than the endothelial barrier. Therefore, the structural integrity of the lung epithelium is vital for normal host function as well as withstanding challenges that could compromise lung function. The manner in which the epithelium provides mechanical barrier protection is multifaceted, involving multiple junctional complexes that include the tight junction, the adherens junction, and desmosomes. With few exceptions, the junctional complexes are located near the apex of all lung epithelial cells that line the airways and alveoli of the lung. The composition of these complexes and formation of cell-to-cell contact involves multiple proteins that are dynamically regulated during normal homeostasis. We predict that during inflammatory stress and release of cytokines, which activate the extrinsic apoptosis pathway, junction complex proteins of the lung epithelium become substrates for terminal caspases, leading to degradation of the junction complex and loss of cell-to-cell contact.
Acute respiratory distress syndrome (ARDS) involves severe acute respiratory failure and is associated with profound loss of epithelial mechanical barrier function (2). Frank injury and loss of the epithelium is a constitutional event that occurs at the onset of ARDS and includes apoptosis of epithelial cells resulting in barrier dysfunction and increased permeability (5, 21, 26). The investigations of the pathophysiological events underlying ARDS, a constellation generally referred to as acute lung injury (ALI), are substantial. Although important discoveries have emerged, the goal of translating these findings into meaningful interventions that reduce morbidity and mortality remains incomplete and elusive (25). The genetic, cellular, molecular and iatrogenic factors that contribute to ALI remain largely unknown. Therefore, further investigation focused on epithelial susceptibility to apoptosis, along with the basic mechanisms that regulate cellular and barrier function, is warranted.
Hypozincemia and shunting of zinc into the cellular compartment are hallmark features of patients who encounter systemic acute inflammation and infection (27). We recently reported (4) findings similar to those of others (9, 30) that zinc acts as a cytoprotectant of the lung epithelium during inflammatory stress and that cellular depletion enhances susceptibility to extrinsic apoptosis. We have also observed (4) that cellular zinc uptake rapidly activates the phosphatidylinositol 3-kinase/Akt signal transduction pathway and protects the epithelium, whereas zinc depletion suppresses both pathways, thereby promoting cell death. In light of these observations, we hypothesize that zinc plays a vital role in regulating epithelial barrier function during inflammatory stress.
The purpose of this investigation was to further evaluate the cytoprotectant role of zinc in human lung epithelia and further elucidate mechanisms, at the onset of ALI, responsible for lung epithelial dysfunction and loss of mechanical barrier function. To accomplish this we focused our attention on β-catenin and E-cadherin, two proteins directly involved in cell-to-cell contact through formation of the adherens junction complex (14, 15). We report on the relationship between caspase-3, a major effector caspase involved in proteolysis during apoptosis, and proteins that form the adherens junction complex. In support of our hypothesis, we observed that zinc deprivation dramatically accelerates caspase-3 activation, in conjunction with inflammatory stress, leading to proteolysis of junction proteins and increased paracellular transit. We also observed that zinc supplementation prevented caspase-mediated events and maintained cellular order. In direct comparison, blockade with a caspase inhibitor inhibited cellular apoptosis but was inferior in preventing barrier dysfunction. We believe 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 reveal molecular insights for alternative therapeutic strategies to prevent ALI.
MATERIALS AND METHODS
Primary cell culture.
Primary human lung upper airway epithelial cells (HUAECs) were isolated after enzymatic dissociation from the 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 (20). Results in this investigation were derived from six different donors obtained during the course of this investigation. In all experiments primary cultures were a minimum of 2 wk past initial seeding. HUAECs were maintained in a 1:1 mixture of Dulbecco's modified Eagle's medium and Ham's F-12 medium (DMEM-F-12) supplemented with 2% Ultroser G (BioSepra; Villeneuve, La Garenne, France) and antibiotics unless otherwise stated. Primary human lung alveolar epithelial cells (HAlvECs) were isolated after enzymatic dissociation of 50 g of parenchymal tissue with a positive selection procedure using magnetic beads combined with antibodies that select alveolar epithelial cells as previously reported (13). We routinely achieve purity >95% with yields >1 × 107 cells per 50 g of donor tissue, which translates into approximately two to three 24-well plates per specimen. The alveolar cells were maintained for ∼2 wk as a differentiated, mixed type I/II culture grown at an air-liquid interface with transepithelial resistance (TEER) recordings that typically exceed 800 Ω. Human lungs were collected with approval from The Ohio State University Institutional Review Board. In preparation of all experiments, unless indicated, the cells were cultured under serum-free conditions for 48 h followed by the addition of 250 U/ml interferon (IFN)-γ (catalog no. PHC 4031, BioSource, Camarillo, CA), 100 ng/ml tumor necrosis factor (TNF)-α (Knoll Pharmaceuticals, Whippany, NY), and/or 200 ng/ml anti-human CD95 (FasAb) (catalog no. MAB 142, R&D Systems, Minneapolis, MN) for an additional 48 h. To achieve zinc deprivation, cultures were grown in non-zinc-supplemented DMEM 48 h before cytokine addition. To deplete intracellular zinc stores, the zinc chelator N,N,N′,N′-tetrakis(2-pyridylmethyl)ethylenediamine (TPEN; 20 μM) (8) was added 4 h before the end of the 48-h incubation period with cytokines. The caspase-3 inhibitor Asp-Glu-Val-Asp-fluoromethylketone (DEVD-fmk) was added to primary cultures immediately before TPEN, unless otherwise stated, at a concentration of 50 μM. All treatment conditions were done in triplicate, and each data set presented is representative of at least three separate experiments unless otherwise stated.
Western blot analysis of HUAEC lysates.
Cell lysis buffer (Cell Signaling, Beverly, MA) with 1 mM PMSF was directly added to the apical surface of the Transwell chamber after removal of the culture medium from the basolateral chamber. Cells were scraped and collected with lysis buffer and then centrifuged at 14,000 rpm for 12 min at 4°C. Supernatants were quantified by protein assay (Bio-Rad, Hercules, CA) and then mixed with Laemmli buffer (Bio-Rad) containing 5% (vol/vol) 2-mercaptoethanol, boiled for 5 min, separated on 10% or 12% SDS-PAGE gel (Bio-Rad), and transferred to a nitrocellulose membrane (Amersham Biosciences, Little Chalfont, UK). Membranes were blocked with 5% (wt/vol) nonfat milk in phosphate-buffered saline (PBS)-0.1% Tween 20 (PBS/T) for 1 h at room temperature and then incubated with primary antibody overnight at 4°C. After washing, the membranes were incubated with the secondary antibody for 1 h at room temperature. Signal was detected with an ECL kit (Amersham Biosciences) and a Fluor-S Multi-Imager Max/Bio-quantity one (Bio-Rad). The following antibodies were used in our experiments: anti-E-cadherin (1:2,500; BD Biosciences, San Jose, CA); anti-β-catenin (1:500; BD Biosciences); anti-caspase-3 (1:1,000; Cell Signaling, Beverly, MA); anti-actin (1:5,000; MP Biomedicals); goat anti-mouse IgG-horseradish peroxidase (HRP) (1:3,000; Cell Signaling); and goat anti-rabbit IgG-HRP (1:3,000; Zymed, San Francisco, CA).
Measurement of caspase activity.
The presence of active caspases was determined by 7-amido-4-trifluoromethylcoumarin (AFC) assay using specific fluorosubstrates. For all AFC preparations, cells (3 × 106) were collected by centrifugation and washed with potassium phosphate buffer containing magnesium sulfate and lysed by four cycles of freeze-thawing as previously described (10). Lysates were incubated with cyto-buffer (10% glycerol, 50 mM PIPES, pH 7.0, 1 mM EDTA) containing 1 mM DTT and 20 μM DEVD-AFC (Enzyme Systems Products). The release of free AFC was determined with a Cytofluor 4000 fluorimeter (Perseptive, Framingham, MA; filters: excitation 400 nm, emission 505 nm).
Analysis of apoptosis.
Cells were detached with a nonenzymatic cell dissociation solution (Sigma, St. Louis, MO) and pooled with cells already suspended during culture followed by cytopsin preparation onto silane-treated slides. Cells were then washed and stained with the M30 CytoDEATH-FITC antibody (Boehringer Mannheim, Indianapolis, IN), a monoclonal antibody that specifically detects caspase-cleaved human cytokeratin-18 (CK-18), as we have previously reported (10). Cells were also stained with 0.5 mg/ml 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI; Roche Molecular Biochemicals, Indianapolis, IN). Specificity was confirmed by comparison against an antibody that recognizes native CK-18 in all cells as well as comparison to an isotype control antibody, MOPC21 (Sigma; not shown). Apoptotic (M30-positive cells) and total (DAPI-stained nuclei) cells were enumerated by a blinded observer who randomly selected six fields of view per treatment condition. In our experience, identical results are obtained with the use of the TUNEL assay, thereby confirming the validity of the M30 assay (10). Data are presented as the average percentage of apoptotic cells divided by the total number of cells per viewing area. Results are compared to actinomycin D-treated cells as a positive control where indicated. Lactate dehydrogenase release was used as an additional measure of cell death in compliance with the manufacturer's instructions (Roche Applied Sciences, Indianapolis, IN).
Assessment of paracellular transport.
TEER was monitored with a portable ohmmeter (Millicell-ERS, Millipore, Bedford, MA). To measure TEER, 300 μl of medium was 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 (3). 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 is a Lucifer yellow flux <0.25%/h. Experimental conditions were compared to 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 occurs within 20 min after the addition of EGTA.
Assessment of apoptotic and necrotic cells.
Apoptotic and necrotic cells were concomitantly analyzed with the YO-PRO staining kit (Vybrant Apoptosis assay Kit no. 4, Molecular Probes, Eugene, OR). Briefly, after treatment, both floating cells and attached cells were collected, pooled, and washed once with PBS and then incubated with 20 nM YO-PRO-1 dye and 1.5 μM propidium iodide in PBS for 30 min at room temperature. The cells were centrifuged by cytospin onto slides after washing with PBS and mounted under coverslips with FluroMount-G (Southern Biotech, Birmingham, AL). The slides were immediately evaluated by fluorescence microscopy. Apoptotic cells stain with moderate green fluorescence, and necrotic cells stain with red fluorescence. Six pairs of images of each slide were randomly evaluated.
Immunodetection of active caspase-3.
Detached and attached cells were collected and washed with PBS and then centrifuged on cytospin slides. After fixation with 4% paraformaldehyde in PBS for 60 min at 4°C, slides were rinsed three times with Tris-buffered saline (TBS)-0.1% Triton X-100 (TBS/T). Nonspecific binding was quenched by incubation in 5% goat serum in PBS/T for 1 h at room temperature. Primary antibody [anti-caspase-3 (1:50) or anti-cleaved caspase-3 (1:200), Cell Signaling] was then applied in 5% BSA in TBS and incubated overnight at 4°C. After being washed three times with TBS/T, slides were incubated with goat anti-rabbit biotin-conjugated antibody (Sigma) for 60 min at room temperature. Slides were washed three times with TBS/T and once with TBS and then incubated with 0.6% H2O2 at room temperature for 30 min. After washing, slides were incubated with avidin-biotin complex reagent (Dako) for 60 min. Slides were incubated with fresh diaminobenzidine solution (Dako) for 5 min, rinsed with distilled water, and counterstained with hematoxylin. After dehydration with alcohols and xylene, slides were mounted on a permanent coverslip and evaluated by light microscopy.
Immunofluorescence stain of E-cadherin.
Primary lung epithelial cells cultured on inserts [treated with or without combination of IFN-γ, TNF-α, and Fas cross-linking antibody (ITF) ± TPEN ± Zn] were washed with PBS on the basolateral side and fixed with 2% paraformaldehyde for 1 h at room temperature. Inserts were then washed with PBS three times, incubated with 5% goat serum in incubation buffer containing 1% BSA and 0.3% Triton X-100 for 1 h at room temperature, and then incubated with anti-E-cadherin (1:100) at 4°C overnight. After washing, cells were incubated with Alexa Fluor 488 goat anti-mouse IgG (Molecular Probes) for 1 h and counterstained with DAPI for 10 min at room temperature. Insert membranes were then cut and mounted on slides, followed by washing. Images were obtained with an Olympus BX61 disk-scanning confocal microscope equipped with a Hamamatsu charge-coupled device camera and SlideBook 2D/3D time lapse imaging software.
Transmission electron microscopy.
Alveolar epithelial cells grown on inserts were fixed with 2% paraformaldehyde and 2% glutaraldehyde in 0.1 M phosphate buffer overnight at 4°C. After fixation the inserts were rinsed several times with PBS, followed by postfixation with 1% osmium tetroxide in phosphate buffer for 1 h. The inserts were dehydrated through a series of graded ethyl alcohols from 70% to 100%. After dehydration, inserts were incubated with two changes of 100% propylene oxide (PO) for 15 min each and finally in a 50:50 mixture of PO and embedding resin for 12–18 h. The specimens were then transferred to fresh 100% embedding medium for at least 1 h, embedded in a fresh change of 100% embedding medium, and incubated for 12–18 h at 60°C for polymerization. The resin blocks were thin sectioned with a diamond knife at 70–90 nm, and sections were then placed on copper mesh grids. After drying on filter paper for a minimum of 1 h, the sections were stained with the heavy metals uranyl acetate and lead citrate for contrast and then viewed by transmission electron microscopy in conjunction with the Ohio State University Microscopy Facility.
All data are expressed as means ± SE. Paired t-tests were used for single comparisons (Microsoft Excel; Microsoft, Redmond, WA). For comparisons that involved multiple variables and observations, two- and three-way ANOVA (JMP; SAS Institute, Cary, NC) were used. Having passed statistical significance by ANOVA, individual comparisons were made with the Tukey multiple-comparison test. Statistical significance was defined as a P value <0.05.
Proinflammatory cytokines and zinc depletion induce caspase-3, apoptosis, and barrier dysfunction.
We previously reported (4) that a combination of IFN-γ, TNF-α, and a Fas cross-linking antibody, referred to here as ITF, did not substantially induce apoptosis or mechanical dysfunction of fully differentiated primary human upper airway lung epithelia. However, ITF in conjunction with zinc deficiency led to substantial apoptosis and increased paracellular transit across the monolayer. To further characterize this, we first exposed primary upper airway epithelial cultures to individual cytokines, in conjunction with the zinc chelator TPEN, and measured apoptosis. We observed a modest increase in apoptosis with TNF-α, IFN-γ, or FasAb in tandem with TPEN treatment; however, we observed that individual cytokine exposure in conjunction with TPEN was less lethal compared with a combination of all cytokines combined with TPEN (Fig. 1A). A dose titration with TPEN was then conducted in ITF-exposed cultures, and we observed a significant increase in apoptosis with higher doses of TPEN, whereas exposure to the highest dose of TPEN resulted in significantly less cell death (Fig. 1B). Primary cultures were then subjected to ITF as previously described and also exposed to TPEN at increasing intervals of time between 0 and 8 h. Exposure to ITF or TPEN alone led to a modest increase in apoptosis above baseline (Fig. 2A). In contrast, increased exposure to TPEN in conjunction with ITF resulted in a substantial increase in the number of apoptotic cells, achieving a maximum of ∼90% cell death at 8 h. Both adherent and disadhered M30-positive cells were analyzed. Interestingly, caspase-3 activity achieved a maximum at 4 h during TPEN exposure and then declined at the later time points (Fig. 2B). Similar results were obtained when caspase-8 and caspase-9 activity were measured in the same samples (data not shown). As expected, increased apoptosis correlated with a decrease in TEER, recorded as Δ resistance (Fig. 2C), and an increase in paracellular flux of Lucifer yellow across the monolayer (Fig. 2D). To confirm caspase-3 activation, we also immunostained epithelia with an antibody that recognizes active caspase-3. Primary cultures exposed to a combination of cytokines and TPEN demonstrated very intense immunostaining throughout the cytoplasm within 4 h after zinc chelation (Fig. 3C), whereas untreated (Fig. 3A) or ITF-treated (Fig. 3B) cultures exhibited minimal presence of active caspase-3.
Zinc depletion alters alveolar epithelium susceptibility to extrinsic apoptosis.
Having observed that primary cultures of differentiated upper airway epithelium are vulnerable to apoptosis when zinc depleted, we wanted to determine whether the same applies to primary HAlvEC cultures. Our rationale was that although ALI affects both the conducting and terminal airway in ARDS the alveolar epithelium is where the major lesion occurs. Before each experiment, freshly isolated type II alveolar cells were allowed to achieve monolayer status with electrically tight junctions at an air-liquid interface. A panel of antibodies and lectins was used to confirm cell phenotype, as previously reported by others (11), before conducting studies (data not shown). Similar to upper airway cultures, HAlvECs were more vulnerable to apoptosis when exposed to ITF in conjunction with the zinc-chelating agent TPEN (Fig. 4A). We also observed that this rapidly led to a compromise in barrier integrity as measured by TEER (Fig. 4B) and Lucifer yellow flux across the monolayer (Fig. 4C). Most striking, we once again observed that zinc supplementation prevented apoptosis and preserved barrier function. An electron micrograph image of a typical HAlvEC culture before treatment is provided in Fig. 4D to demonstrate morphological evidence of a uniform monolayer with cell-to-cell contact.
Zinc depletion enhances degradation of adherens junction complex.
Having shown that zinc depletion enhances caspase-3 activation and programmed cell death and leads to increased permeability across the monolayer, we turned our attention to β-catenin and E-cadherin. Both proteins work in concert to establish cell-to-cell contact at the adherens junction complex (14, 15) and are also known substrates for caspase-3 (6, 28, 29). Again, primary upper airway epithelia were exposed to ITF and zinc depletion, and then whole cell lysates were analyzed by Western blot analysis. We observed that the combination of ITF plus zinc depletion resulted in the rapid conversion of both β-catenin and E-cadherin from their respective molecular masses of 120 and 92 kDa to caspase-cleaved degradation products (Fig. 5, A and B). This occurred in parallel with conversion of caspase-3 from its inactive to its active form (Fig. 5C) and in tandem with a precipitous drop in TEER and an increase in permeability across the monolayer. In comparison, ITF or TPEN treatment alone did not induce caspase-mediated events. Zinc supplementation prevented ITF+TPEN caspase-mediated degradation of the junction proteins. Changes in TEER and permeability are not shown in Fig. 5 since identical findings were obtained compared with those previously presented (see Fig. 2, C and D).
In addition to determining that zinc is an essential cytoprotectant required by the lung epithelium, we evaluated its potential as a therapeutic agent to protect against cell death and mechanical dysfunction. Since caspase-3 was identified as an important mediator in our model, we also evaluated the therapeutic potential of the caspase-3 inhibitor DEVD-fmk to prevent acute lung injury. Similar to past experiments, we exposed cultures to ITF for 48 h, followed by the addition of TPEN during the last 4 h. Cultures were supplemented with zinc or DEVD-fmk at the same time as TPEN or at 1- and 2-h intervals afterward. The effect of zinc or DEVD-fmk treatment was examined by measuring apoptosis, caspase activity, and paracellular transit. We observed that both zinc and DEVD-fmk were effective at inhibiting caspase activation (Fig. 6A) and apoptosis (Fig. 6B), even when administered up to 2 h after TPEN. However, only zinc supplementation was effective at preserving barrier function by maintaining barrier function across the monolayer, whereas the caspase inhibitor was not as effective at preventing transit of Lucifer yellow across the monolayer (Fig. 6C). In addition, zinc was more effective at preventing apoptosis in comparison to DEVD-fmk (Fig. 6D). Further evaluation of HUAECs revealed that only zinc supplementation prevented degradation of β-catenin (Fig. 7A) and E-cadherin (Fig. 7B), whereas the caspase-3 inhibitor did not. In contrast to previous measurements of caspase enzyme activity, DEVD-fmk also failed to inhibit the conversion of caspase-3 to the active form as detected by Western blot analysis (Fig. 7C). These observations were further supported by visualizing E-cadherin in HUAECs grown on Transwell inserts.
Immunostaining of fully differentiated HUAEC cultures demonstrated a predictable pattern of E-cadherin localization at the cell membrane, particularly at points of cell-to-cell contact (Fig. 8A). After exposure to ITF and zinc depletion, the continuity of E-cadherin staining is disrupted with evidence of cellular redistribution in the cytoplasm concomitant with increased disorder across the monolayer as shown in Fig. 8B, inset. Zinc supplementation, to a large extent, was able to maintain a normal pattern of E-cadherin staining at cell junctions and preserve monolayer integrity (Fig. 8C), whereas treatment with the caspase inhibitor did not (Fig. 8D).
As a last measure we determined whether the disparity in preserving barrier function between zinc and the caspase inhibitor was attributable to differences in apoptosis and necrosis. To accomplish this we utilized the YO-PRO-1 dual-staining kit, which detects apoptosis and necrosis on the basis of changes that occur in the permeability of cell membranes. YO-PRO-1 stain selectively passes through the plasma membranes of apoptotic cells and labels them with moderate green fluorescence, whereas necrotic cells are detected by red fluorescence with propidium iodide. In support of our previous findings, we observed that ITF+TPEN primarily induced apoptosis compared with untreated cells, which showed little evidence of apoptotic or necrotic cells (Fig. 9). Zinc supplementation substantially inhibited the number of apoptotic-appearing cells, with infrequent occurrence of necrotic cells. In sharp contrast, although treatment with DEVD-fmk suppressed the number of apoptotic cells, a substantial increase in necrotic cells was observed.
We previously reported (4) that zinc deficiency predisposes differentiated cultures of primary human upper airway epithelia to inflammation-mediated apoptosis. To more carefully characterize the relationship between the lung epithelium, zinc, and ALI, we expanded our studies, using primary differentiated cultures of human upper airway as well as alveolar epithelia. Our initial observations revealed that the lung epithelium becomes increasingly vulnerable when exposed to multiple cytokines, as opposed to individual cytokines, that activate death receptor-mediated apoptosis, but only in the presence of intracellular zinc deficiency. The magnitude of cell death and barrier dysfunction was directly proportional to the degree of intracellular zinc depletion in addition to the time of exposure to zinc depletion in conjunction with acute inflammation. Furthermore, the effects of zinc depletion, cellular demise, and barrier dysfunction were directly linked to caspase-3 activation and proteolysis of key proteins that establish cell-to-cell contact. Zinc supplementation was highly effective at preserving cell integrity and barrier function, whereas the caspase-3 inhibitor DEVD-fmk was less effective and did not prevent loss of barrier function.
We believe that our findings have implications for lung epithelial function and the host response during acute inflammation. Hypoferremia and hypozincemia are two of the many changes that occur in humans during the initial stages of the acute-phase response (27). Hypoferremia is associated with an innate host defense response to decrease the availability of this essential metal to pathogenic organisms (18). Hypozincemia results from zinc redistribution into the cellular compartment, although the rationale to account for this is much less understood. In vitro studies utilizing human peripheral blood mononuclear cells indicate that cellular zinc uptake acts as an immunostimulant through enhanced cytokine production (33, 34) and support human studies demonstrating enhancement of immune function following zinc supplementation (1, 7). In our investigation we consistently observed that zinc has a cytoprotective role during acute inflammation. In particular, zinc depletion significantly increases vulnerability to extrinsic apoptosis, which is in sharp contrast to acute intracellular zinc repletion, which prevents cell death. Our interpretation of these findings is that zinc mobilization into the lung epithelium, at the onset of the acute-phase response, is an essential innate host protective response that prevents cell death. With respect to previous work that identified zinc as an immunostimulant, we propose that a dual, complementary role for zinc may exist during host defense whereby zinc enhances host immune function while simultaneously protecting nonimmune cells. Our results also suggest that abnormal zinc homeostasis, due to either dietary insufficiency or genetic variability, may increase host susceptibility to cell damage and tissue dysfunction, as is the case in patients who suffer from acrodermatitis enteropathica (AE) (32). Although AE is considered a rare disease, it is also recognized that zinc deficiency is underestimated and affects millions of individuals in the United States, thereby suggesting that these findings have relevance to a much larger population (31).
The probability of genetic variability as a cause for tissue dysfunction is quite high when considering the complexity of zinc homeostasis in humans. Zinc metabolism is regulated by zinc transporters encoded by two solute-linked carrier (SLC) gene families: SLC30/ZnT and SLC39/Zip. Our preliminary evaluation of the human genome (http://genome.ucsc.edu) revealed 10 SLC30 and 14 SLC39 transporters, corroborating previous reports (12, 22). The two families have opposite roles in cellular zinc homeostasis. SLC30 transporters reduce intracellular zinc availability by promoting zinc efflux from cells, whereas SLC39 transporters increase intracellular zinc availability by promoting cytosolic zinc uptake. SLC30 and SLC39 transporters also exhibit unique tissue specific expression and function (19, 22). In particular, expression of the murine zinc transporter ZIP14 increased substantially compared with other zinc transporters after injection with LPS or turpentine, factors used to invoke systemic inflammation and the acute phase response in mice (23). In particular, ZIP14 protein was upregulated, localized at the cell membrane, and resulted in rapid mobilization of zinc into cells. Furthermore, these events required transcriptional activation by interleukin-6, a prototypical inflammatory cytokine associated with ARDS. In light of these findings it is interesting to speculate that specific human zinc transporters may have an important role in regulating zinc homeostasis during inflammation in order to protect the host.
Further evaluation of zinc as a therapeutic agent to prevent ALI is warranted especially if at-risk populations can be identified. Since alveolar epithelial injury is a strong predictor of ARDS patient outcome (24, 35) and the alveolus is accessible to topical drug delivery, the prospect of aerosol-based zinc delivery to the lung to prevent epithelial injury and dysfunction is appealing. Topical administration is also attractive when considering the rapid onset of ALI, the narrow window for therapeutic intervention, and our observation that zinc supplementation sustains lung epithelial barrier function even when administered within hours after the onset of inflammation. In comparison to the caspase-3 inhibitor DEVD-fmk, zinc rescue was superior in that it prevented caspase activation and apoptosis and fully maintained barrier integrity. At face value, the caspase inhibitor also appeared to be beneficial in that it prevented apoptosis; however, barrier function further deteriorated. Similar effects were observed despite efforts to lower the dose of DEVD-fmk (data not shown). Furthermore, DEVD-fmk did not appear to completely prevent caspase activation, perhaps due to an inability of the compound to access all pools of active enzyme. On further evaluation, we observed that barrier dysfunction was caused by a substantial shift to necrosis, perhaps indicating that inhibition of apoptosis at a terminal end point, although preventing apoptosis, is too late to prevent cell death. This type of phenomenon was previously identified with this class of compound (17). On the basis of our observations, we conclude that pharmacological strategies to inhibit terminal caspase activation may have limitations if the intended drug target is the lung epithelia.
Our investigation provides new evidence on the relationship between zinc, human airway and alveolar epithelial cells, and regulation of the epithelial response to inflammatory stimuli involved in lung pathogenesis. We observed that zinc deprivation substantially increases susceptibility to extrinsic apoptosis and facilitates barrier disruption in both epithelial populations. This supports the concept that host predisposition to ALI is determined by inflammation in addition to cofactors that regulate the cellular response to inflammatory stress. In light of this, we predict that dietary zinc intake as well as genetic factors that regulate intracellular zinc stores are vital components of lung epithelial cell homeostasis and deserve further study. The supportive contribution of zinc to innate immune defense in the lung has great appeal in terms of helping to 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 the National Heart, Lung, and Blood Institute (HL-56336, D. L. Knoell) and The American College of Clinical Pharmacy (Career Investigator Award, D. L. Knoell).
We express special thanks to Lifeline of Ohio Tissue Procurement Agency.
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