Department of Medicine, University of Adelaide, The Queen Elizabeth Hospital, Woodville, South Australia 5011, Australia
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The respiratory
epithelium is vulnerable to noxious substances, resulting in the
shedding of cells and decreased protection. Zinc (Zn), an antioxidant
and cytoprotectant, can suppress apoptosis in a variety of cells. Here
we used the novel Zn-specific fluorophore Zinquin to visualize and
quantify labile intracellular Zn in respiratory epithelial cells.
Zinquin fluorescence in isolated ciliated tracheobronchial epithelial
cells and intact epithelium from sheep and pigs revealed an intense
fluorescence in the apical and mitochondria-rich cytoplasm below the
cilia. Zinquin fluorescence was quenched by the Zn chelator N,N,N',N'-tetrakis(2-pyridylmethyl)ethylenediamine
(TPEN) and increased by the Zn ionophore pyrithione. We also assessed
whether changes in intracellular labile Zn would influence
susceptibility of these cells to apoptosis by hydrogen peroxide. Our
results confirm that Zn deficiency enhanced hydrogen peroxide-induced caspase activation from 1.24 ± 0.12 to 2.58 ± 0.53 units · µg protein1 · h
1
(P
0.05); Zn supplementation suppressed these effects.
These findings are consistent with the hypothesis that Zn protects
upper respiratory epithelial cells and may have implications for human asthma where there is hypozincemia and epithelial damage.
respiratory epithelium; Zinquin
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
DAMAGE TO THE
AIRWAY EPITHELIUM by inhaled noxious agents (e.g., environmental
pollutants) not only compromises its protective barrier function but
also results in decreased production of smooth muscle relaxant factor
and increased release of proinflammatory cytokines such as
interleukin-6, interleukin-8, granulocyte-macrophage colony-stimulating
factor, and tumor necrosis factor- (22). Although the
mechanism of respiratory epithelial cell death has not been extensively
studied, in at least two reports (3, 33), including an in vitro model of tracheal regeneration, it was shown that
the death of these cells was due to apoptosis. Another example where
apoptosis of these cells may be important is in asthma, where there is
an increase in the fragility and shedding of the columnar airway
epithelium (19). In these cases, apoptosis may contribute
to the denuding of the respiratory epithelium and increased exposure of the basement membrane to toxic mediators of inflammatory cells, thereby exacerbating inflammation while retarding airway repair
(3, 13, 19, 27).
One factor that may be significant in the biology of the respiratory epithelium is the group IIb metal zinc (Zn), an important antioxidant and anti-inflammatory agent that participates in various cellular events and is a cofactor for many metalloenzymes and transcription factors (32). Intracellular Zn exists in two discreet pools, the first of which is nonexchangeable and tightly bound to metalloenzymes and the second is a more labile and dynamic pool that is rapidly exchangeable and able to be altered by Zn deprivation or supplementation (35). One important role of labile Zn is as a survival factor that suppresses apoptosis. Studies from our laboratory (35) and others (12, 15, 16) have shown that Zn deficiency in vitro and in vivo stimulates internucleosomal DNA fragmentation and apoptosis in lymphoid and myeloid cells as well as in intestinal epithelium, neural epithelium, and endothelium. Zalewski et al. (35) have demonstrated an inverse correlation between the level of intracellular labile Zn in lymphocytes and their susceptibility to DNA fragmentation. These results imply that a reduction below a threshold concentration in this Zn pool facilitates apoptotic DNA fragmentation.
Zn is important for the integrity, growth, and repair of epithelial tissues in the skin and gastrointestinal tract (1, 12), but its role in the respiratory epithelium has not yet been studied. We propose that intracellular labile Zn is important in this tissue as a cytoprotectant against toxins and inflammatory mediators in a way similar to that reported for the endothelium (16). Therefore, the aim of this study was to investigate where Zn is localized along and within the respiratory epithelium and to further elucidate its role in regulating apoptosis of these cells. For this study, we used a variety of cells and tissues that included 1) NCI-H292 and A549 cells (malignant cell lines representative of bronchial and alveolar epithelia, respectively), 2) primary pig and sheep ciliated epithelial cells, and 3) cryostat tissues from these animals.
To assess the levels and distribution of intracellular labile Zn, we used the ultraviolet (UV)-excitable Zn-specific fluorophore Zinquin that has previously enabled us to visualize and image distinct pools of labile Zn in a range of cell types and tissues (6, 35-37). The Zn chelator N,N,N',N'-tetrakis(2-pyridylmethyl)ethylenediamine (TPEN) and the Zn ionophore sodium pyrithione (35) were used to determine the effects of changes in the intracellular Zn status of these cells on their susceptibility to undergo apoptosis induced by the oxyradical hydrogen peroxide (H2O2) (25) or the short chain fatty acid butyrate (24).
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Materials
The major materials were EDTA, EGTA, herring sperm DNA, Hoechst 33258, Nonidet P-40 (NP-40), dithiothreitol, sucrose, HEPES, TPEN, sodium pyrithione, amphotericin B, insulin, transferrin, epidermal growth factor (EGF), protease (from Streptomyces griseus), and retinoic acid (all from Sigma, St. Louis, MO); hematoxylin staining solution, eosin staining solution, H2O2 (30% wt/vol), and sodium butyrate (all from BDH, Poole, UK); LHC-9 basal medium (Clonetics, Walkersville, MD), RPMI 1640 medium, penicillin-streptomycin, EDTA-trypsin, L-glutamine, 3-[(3-cholamidopropyldimethylammonio)-1-propanesulfonate] (CHAPS; ICN, Aurora, OH); fetal bovine serum (Biosciences, Sydney, Australia), Percoll (Amersham Pharmacia Biotech, Uppsala, Sweden); gentamicin (David Bull Laboratories, Melbourne, Australia); Asp-Glu-Val-Asp 7-amino-4-(trifluoromethyl)coumarin (DEVD-AFC) and Tyr-Val-Ala-Asp-AFC (YVAD-AFC; both from Kamiya Biomedical, Tukwila, WA); Val-Asp-Val-Ala-Asp-AFC (VDVAD-AFC), Leu-Glu-Val-Asp-AFC (LEVD-AFC), Trp-Glu-His-Asp-AFC (WEHD-AFC), Val-Glu-Ile-Asp-AFC (VEID-AFC), and Leu-Glu-His-Asp-AFC (LEHD-AFC; all from Calbiochem); epoxypropane and Procure 812 resin (both from Electron Microscopy Sciences, Fort Washington, PA); Tissue-Tek optimum cutting temperature compound (Miles); and ethyl-(2-methyl-8-p-toluenesulfonamido-6-quinolyloxy)acetate [Zinquin (21); Department of Chemistry, University of Adelaide, Adelaide, Australia]. All other reagents not listed were of reagent grade unless otherwise indicated.Cell Lines
A549 and NCI-H292 cells were obtained from Dr. D. Knight (Queen Elizabeth II Medical Center, University of Western Australia, Nedlands, Australia) and cultured in RPMI 1640 medium (24). A549 cells were derived from a human alveolar cell carcinoma; they are epithelial in morphology and hyperdiploid and represent type II alveolar cells (20). NCI-H292 cells were derived from a cervical node metastasis of a pulmonary mucoepidermoid carcinoma; they are representative of the upper bronchial respiratory airways and are epithelial in morphology, containing numerous mucin-secreting granules (5).Respiratory Epithelial Tissues and Tracheobronchial Cell Isolation
Pig and sheep respiratory tracts were removed from freshly killed animals and rinsed in chilled phosphate- buffered saline solution (pH 7.4). Samples were frozen in optimum cutting temperature compound and liquid nitrogen for cryostat sections or fixed in Formalin and embedded in paraffin for histological assessment. Cryostat sections were stained with hematoxylin and eosin for morphological assessment. Collagen was identified with Weigert's hematoxylin and van Gieson's stain (a gift from the Histopathology Department, University of South Australia, Adelaide, Australia). These experiments conformed to the National Health and Medical Research Council of Australia animal ethics guidelines.To prepare isolated epithelial cells, excised segments were incubated at 4°C overnight in RPMI 1640 medium containing 0.05% protease. Dissociated cells were removed by vigorous agitation for 5 min, and enzymatic digestion was terminated with fetal bovine serum (10% final concentration). Cells were centrifuged at 150 g for 5 min to produce a cellular pellet containing isolated mucus cells, ciliated cells, and nonciliated basal cells that were used for Zinquin fluorescence studies. To prepare a highly enriched population of ciliated cells for apoptosis assays, the Percoll separation technique of Takizawa et al. (31) was used. Cell viability as determined by trypan blue exclusion was >90%. The upper band contained a highly enriched population of ciliated cells that were used for Zn and apoptosis studies. Epithelial cells were cultured in a modified serum-free LHC-9 medium supplemented with 2 mM L-glutamine, 25 µM amphotericin B, 1 µg/ml of insulin, 1 µg/ml of transferrin, 10 ng/ml of EGF, and 10 ng/ml of retinoic acid. Where extensive clumping had occurred, cell suspensions were passed two times through a 20-gauge needle before being seeded into wells.
Depletion and Augmentation of Intracellular Labile Zn
To deplete intracellular Zn, the cells were incubated with varying concentrations (up to 25 µM) of TPEN for 1 h at 37°C in complete culture medium. TPEN was stored as a 5 mM stock solution in DMSO atQuantification of Zinquin Fluorescence by Image Analysis
Cryostat sections 3 µm thick from animal airways and alveolar tissue were fixed in acetone for 5 min at room temperature (RT) and then rinsed in PBS. Zinquin was freshly diluted in PBS to a final concentration of 25 µM and immediately pipetted onto these sections. After incubation for 30 min at RT in dark and humidified conditions, the sections were mounted with a fluorescent mounting medium (DAKO). An autofluorescence control (PBS alone) was set up for each section to ensure the specificity of Zinquin.A549 and NCI-H292 cells were grown to semiconfluence on sterile glass coverslips in six-well plates for 48 h before the addition of Zinquin. The coverslips were then washed in PBS and incubated with 25 µM Zinquin in PBS for 30 min at 37°C. Primary epithelial cells were incubated with Zinquin (25 µM) as cell suspensions immediately after isolation. In some experiments, intracellular Zn was either depleted or supplemented as in Depletion and Augmentation of Intracellular Labile Zn before the addition of Zinquin.
Fluorescence was examined as previously described (37) with modifications. For cryostat sections, both fluorescence images and corresponding phase-contrast microscope images were captured with a ×20 objective lens and stored. For Zinquin fluorescence in cryostat sections, the outline of the airway epithelium was traced, stored, and copied onto the corresponding fluorescence image. A profile line was then drawn at 90° to the epithelial surface so that it spanned the entire epithelium. The Video Pro image analysis system (Leading Edge) was used to quantify the fluorescence intensities at 1-pixel intervals along the profile line. For each image, seven randomly positioned profiles were obtained, and the mean fluorescence intensity was calculated for each 2.5% distance across the epithelium. As a result, 40 intervals were measured, the first beginning at the luminal surface and the last terminating at the basement membrane. For the alveolar cryostat tissue sections, 20 squares were drawn as overlays within the boundaries of the epithelium on the light images. These overlays were transferred to the corresponding fluorescence image, and the intensities were measured. The mean fluorescence intensity was collated from seven images. For transformed cells, lines were drawn around their borders, and the intensity was quantified. For all images, background fluorescence was determined and subtracted. At least 150 cells or 6-10 tissue sections in each group were analyzed.
For UV laser confocal microscopy, a Bio-Rad MRC-1000 UV laser scanning confocal microscope system equipped with an UV argon laser was used in combination with a Nikon Diaphot 300 inverted microscope in fluorescence mode, with excitation at 363/368 nm and emission at 460 nm with a long-path filter. In the case of ciliated cells, those that were stationary were chosen for z-series imaging of 0.5-µm slices that were 2 µm apart. Images were collected with a ×40 objective water-immersion lens with a numerical aperture of 1.15. Each image was averaged over three scans by Kalman filtering.
Transmission Electron Microscopy of Sheep Tracheal Ciliated Epithelial Cells
Cells (106cells/ml) obtained by protease treatment and Percoll separation were centrifuged at 2,000 rpm, and the pellet was resuspended in 1 ml of an electron microscopy fixative mixture (1.5% glutaraldehyde and 4% formaldehyde in 0.1 M sodium cacodylate buffer) overnight at 4°C. The cell suspension was centrifuged between each of the following processing steps for 1 min at 2,000 rpm. The cell pellet was postfixed in 2% osmium tetroxide and stained en bloc in 2% uranyl acetate. The cells were then dehydrated through a graded series of alcohols (70, 90, and 100%) and further processed through 100% epoxypropane, a 50:50 mixture of epoxypropane and Procure 812 resin (Electron Microscopy Sciences), and two times through 100% Procure 812 resin. The pellet was embedded in polyethylene capsules and cured overnight at 90°C. Survey sections were cut at ~2 µm and stained with toluidine blue. Thin sections were cut at silver-gold color (~100 nm) and mounted on copper grids. The sections were stained with Reynold's lead citrate and examined with a Hitachi H-600 transmission electron microscope at a potential difference of 75 kV.Induction of Apoptosis in Primary and Transformed Cell Cultures
Percoll-separated primary ciliated cells (4 × 104 cells/ml) were cultured as cell suspensions with H2O2 in 24-well plates (NUNCLON) in serum-free modified LHC-9 medium (31). Transformed cells were seeded in six-well plates (Sarstedt, Newton, NC) and allowed to grow to near confluence for 48 h before the addition of apoptotic stimuli (TPEN and/or H2O2 or butyrate). For experiments involving TPEN alone, the cells were incubated with varying concentrations of TPEN (0-50 µM) for 4 h at 37°C. In experiments involving the interaction between TPEN and either H2O2 or butyrate, the cells were initially treated with H2O2 (0-0.125 mM) or butyrate (0-2 mM) for 18 h at 37°C before the addition of TPEN for a further 4 h. For experiments involving the effects of Zn supplementation on apoptosis, sodium pyrithione (1 µM) and ZnSO4 (25 µM) were added 1 h before the addition of the apoptotic inducer.Preparation of Cell Lysates for Caspase Activation Assay
At the end of the incubation period with the apoptotic inducers, the cell lysates were produced. Slight modifications to the methods were made depending on whether the cells were in suspension, e.g., primary cells, or adherent, e.g., transformed cell lines. The primary epithelial cells were removed into tubes and spun at 400 g for 5 min. The cell pellets were then incubated with 500 µl of cell lysis buffer (0.5% NP-40, 5 mM Tris · HCl, and 5 mM EDTA) overnight at 4°C. For A549 and NCI-H292 cells, floating (largely apoptotic) cells were removed first into tubes and centrifuged into a pellet. The remaining adherent cells in the wells were lysed in 750 µl of NP-40 lysis buffer for 15 min at RT. The pellets were recombined with corresponding lysates of adherent cells to yield lysates of total cell population from each well. For all of the cell lysates, insoluble material was pelleted at 12,000 g, and the supernatants were collected and stored atMeasurement of Caspase Activity
Caspase-3-like activity was assayed by cleavage of the fluorogenic substrate Z-DEVD-AFC (as described in Ref. 17). Other substrates used for the detection of various caspase activities in these experiments were YVAD-AFC (caspase-1), VDVAD-AFC (caspase-2), LEVD-AFC (caspase-4), WEHD-AFC (caspase-5), VEID-AFC (caspase-6), and LEHD-AFC (caspase-9). It should be mentioned that although these substrates are preferentially cleaved by the indicated caspases, there may be some overlap. The caspase activities are expressed in fluorescence units per microgram of protein per hour.Other Assays for Apoptosis
Apoptosis was also assessed in some experiments by chromatin condensation in hematoxylin and eosin-stained cell-cultured coverslips and by labeling of nuclear fragments with Hoechst dye 33258 (1 µg/ml). One thousand cells were scored from replicate tubes.Experimental Design
All experiments were repeated a minimum of three times. Typical experiments are described or data were collated as indicated. Significance was determined by Student's t-test. ![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Zinquin Fluorescence Studies
NCI-H292 and A549 cells.
Figure 1 shows corresponding UV laser
confocal light transmission and Zinquin fluorescence images
for NCI-H292 (A and B) and A549 (C and
D) cells, representative of upper airway epithelial cells
and type II alveolar epithelial cells, respectively. NCI-H292 cells had
significantly higher basal levels of Zn as determined by the twofold
increase in Zinquin fluorescence (mean 23.1 ± 2.2 pixels;
n = 29 cells) compared with the A549 cells (mean 11.8 ± 0.3 pixels; n = 192 cells), typical of other labile
Zn-poor cells (35). Zinquin fluorescence was significantly
(P 0.05) quenched by 25 µM TPEN, which has a higher
affinity for Zn than for Zinquin (35). TPEN treatment
reduced fluorescence to 9.7 ± 0.7 pixels in NCI-H292 cells
(n = 28) and 7.2 ± 0.3 pixels in A549 cells
(n = 187). Zn supplementation with exogenous
ZnSO4 and 1 µM sodium pyrithione increased fluorescence
of NCI-H292 cells to 95.1 ± 3.3 pixels (4.1-fold;
n = 58 cells) and to 59.9 ± 1.6 pixels in A549
cells (5.5-fold; n = 307 cells).
|
Isolated tracheal ciliated cells.
Isolated ciliated epithelial cells from the tracheobronchial region of
sheep and pigs were released by proteolytic digestion and purified by
Percoll separation. Figure 2 shows a
transmission electron micrograph of a typical ciliated sheep tracheal
epithelial cell that was used for the following Zinquin fluorescence
and apoptosis studies. These cells, which were viable and had rapidly beating cilia, were incubated with Zinquin in wet suspensions, and
fluorescence was analyzed by UV laser confocal microscopy. Only those
ciliated cells that were not moving across the microscope field were
chosen for z-series confocal images. A bright-field image
and 11 fluorescence images (each from 0.5-µm slices at 2-µm intervals) were captured. Autofluorescence intensity was very low, and
fluorescence gains were set at levels to exclude this.
|
|
Cryostat sections of tracheobronchial and alveolar epithelia. Next, we examined Zinquin fluorescence in frozen sections of sheep and pig tracheobronchial and alveolar tissues. Cryostat sections 3 µm thick were acetone fixed, incubated with and without Zinquin in PBS for 30 min, washed, and mounted for epifluorescence microscopy.
The sections incubated with PBS alone served as controls for UV autofluorescence. Autofluorescence was largely confined to the walls of blood vessels and to the lamina propria where it was colocalized with collagen as detected by staining with Weigert's hematoxylin and van Gieson stain. Collagen fibers were stained bright red, whereas smooth muscle and other cytoplasmic proteins were stained yellow. The stained collagen and the autofluorescence were more structurally organized in the lamina propria sections, particularly in the bronchi and bronchiolar regions of pig, compared with the tracheae or sheep tissues. Importantly, there was no autofluorescence in the epithelia of any of the sections examined. Figure 3, G and H, shows typical hematoxylin and eosin-stained sections and Zinquin fluorescence images, respectively, of frozen sections of pig tracheae. The pseudostratified, ciliated epithelium was lined continuously at the luminal surface (arrow) by a region of intense Zinquin fluorescence. In the other two pigs and in the sheep, there was much less collagen. Importantly, there was no autofluorescence in the epithelium in any of the sections examined. Figure 3I shows another representative section from pig bronchi at a higher magnification and demonstrates the striking demarcation between the labile Zn-rich outer area and the relatively Zn-poor middle to inner layers of the epithelium. Similar data were observed with sheep bronchi and tracheae where there was intense Zinquin fluorescence at the luminal surface (data not shown). The alveolar epithelium in both pig (Fig. 3J) and sheep (data not shown) had a typically dull fluorescence. No distinction in fluorescence was evident between the type I and II epithelial cells. Figure 4 shows a compilation of data from all of the cryostat sections analyzed. The width of the Zinquin fluorescence band varied between different sections from different regions of the track and between sheep and pig specimens. To determine the mean proportion of the epithelium that was labile Zn rich, the epithelium was subdivided into 40 evenly distributed intervals across its entire width, and the mean fluorescence within each interval was determined. For each image, seven randomly placed profile lines (as in Fig. 3H) measuring fluorescence intensity across the epithelium were obtained, and a total of six to eight images from sections of pig and sheep tracheae and bronchioles were quantified. There were insufficient sections of sheep tracheae to do multiple analyses. Similar autofluorescence determinations were made on a separate set of images derived from sections incubated without Zinquin.
|
Caspase Activation and Apoptosis Studies in Zn-Manipulated Cells
The strong staining for Zn in the apical region of airway epithelium coupled with the relatively weak fluorescence in the alveolar epithelium is similar to that reported for procaspase-3 in human tissues by Krajewska et al. (18). These similarities, albeit in different species, in addition to the known suppressive effects of labile Zn on apoptosis, prompted us to investigate whether the labile Zn in respiratory epithelial cells might influence caspase-3 activation and apoptosis.Induction of caspase-3 activation and apoptosis by the Zn chelator
TPEN in primary sheep ciliated epithelial cells and malignant cell
lines.
To determine the effects of lowering intracellular labile Zn on caspase
activation and apoptosis, primary and transformed epithelial cells were
rendered Zn deficient by TPEN. Figure 5, A and B, shows the morphological changes
associated with apoptosis in a typical Zn-depleted primary sheep
ciliated epithelial cell. Fragmentation of the cells into apoptotic
bodies (Fig. 5A), some containing remnants of the nucleus as
shown by the corresponding fluorescence micrographs of chromatin
stained by Hoechst dye 33258 (Fig. 5B), is a common feature
of apoptotic cell death. TPEN also induced DNA fragmentation in these
cells (data not shown).
|
Time course of caspase activation with TPEN. The aim of the following experiment was to determine the optimal time of induction of caspase-3 activity after the decline in intracellular labile Zn induced by 25 µM TPEN. Due to limitations in the number of highly purified primary cells, this time course was performed only in NCI-H292 cells. Experiments were performed in triplicate. To monitor Zinquin fluorescence in the cells, NCI-H292 cells were cultured onto sterile glass coverslips while the caspase-3 activity was determined on the same cells that had grown on the bottom of the six-well culture plates. TPEN was added at time 0 to both populations at varying intervals up to and including 4 h.
Zinquin fluorescence was determined in 17-18 image fields (total of 140-260 cells) for each time point. Mean fluorescence intensity decreased by 38% in the first 30 min after the addition of 25 µM TPEN [from the control value of 23.7 ± 1.3 units · µg protein
|
Suppression of DEVD-caspase activation in Zn-supplemented primary
respiratory epithelial cells.
The previous studies have shown that lowering the intracellular labile
Zn content by TPEN induces DEVD-caspase activation. The effect of
increasing intracellular labile Zn content in primary sheep tracheal
ciliated cells on DEVD-caspase activation was determined next.
DEVD-caspase activation was induced by treatment of the cells overnight
with the apoptosis-inducer H2O2 (0.125 mM).
Intracellular labile Zn was raised by treating the cells with 1 µM
sodium pyrithione plus 25 µM ZnSO4. In cells receiving
supplemented Zn, there was a 59.3% inhibition of
H2O2-induced DEVD-caspase activation (data pooled from 3 experiments, each in triplicate; P 0.005).
Synergstic interactions between TPEN and other inducers of
apoptosis in DEVD-caspase activation.
To determine whether Zn depletion was able to influence toxin-induced
caspase-3 activation, two different classes of apoptosis-inducing agents, H2O2 and butyrate, were used. The
results were collated from at least two experiments each performed in
triplicate for the three different types of cells. Figure
7A shows the data for primary
cells obtained for suboptimal concentrations of TPEN (6.25 µM) and
H2O2 (125 µM) either alone or in combination.
H2O2 alone gave an increase over the control
value for DEVD-caspase activity of 1.24 ± 0.12 units · µg protein1 · h
1;
TPEN alone gave an increase of 0.52 ± 0.14 units · µg
protein
1 · h
1, whereas the two in
combination gave an increase of 2.58 ± 0.53 units · µg protein
1 · h
1.
This increase was significantly greater than the increase that would have occurred if TPEN and H2O2 were
acting additively (P
0.05; Fig. 7A,
dashed line). Similar results were found with NCI-H292 (Fig.
7B) and A549 (Fig. 7C) cells.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
This paper is the first to visualize and quantify labile intracellular Zn in different populations of airway epithelium. There have been no previous studies of Zn distribution in the respiratory system due to lack of sensitive techniques to visualize and quantify tissue Zn. Here, we used the Zn fluorophore Zinquin, which is specific and sensitive for labile intracellular Zn in the nanomolar range (6, 21, 36, 37). Zinquin detects the free or more loosely bound (labile) pools of intracellular Zn. The major findings of this present study are that 1) tracheobronchial epithelial cells are relatively rich in labile Zn compared with alveolar epithelial cells, 2) this Zn is especially concentrated in the mitochondria-rich, apical cytoplasmic region immediately below the cilia, and 3) chelation of Zn with TPEN results in the rapid activation of caspase-3-like activity and downstream events of apoptosis as well as in the markedly enhanced susceptibility of these cells to toxin-induced apoptosis.
Our first finding of labile Zn lining the apical and luminal sides of the entire length of the conducting airways may have an analogy with the concept of Zn in galvanization where Zn acts to protect the underlying steel from oxidation and corrosion. It has been proposed that Zn became particularly important in eukaryotic cell evolution at about the time the atmosphere was becoming oxygen rich and the cells were developing mitochondria (7). Therefore, Zn, as one of the major antioxidants in our body, may play an important role in the protection of the respiratory epithelium. Several properties of Zn contribute to its antioxidant functions. First, unlike Cu and Fe, Zn exists in only one oxidation state (II) and therefore cannot react with oxidants to generate potentially damaging oxyradicals (7); second, Zn may protect cellular membranes, proteins, and DNA from oxidative damage (32); and third, Zn is an important component of the antioxidant Cu/Zn superoxide dismutase (8).
The finding of a low labile Zn content in A549 cells derived from transformed alveolar type II epithelium compared with that in the transformed bronchial epithelial NCI-H292 cells is consistent with the observation of low Zinquin fluorescence in cryostat sections of alveolar epithelium of sheep and pigs. The difference in this fluorescence intensity between the two malignant cell lines cannot be attributed to environmental factors because both cell lines had been maintained for several generations in the same culture medium. Rather, variations in fluorescence may be due to intrinsic differences in the capacities of the two cell types to take Zn up from the medium, perhaps as a consequence of altered levels of Zn transporters in the cell membrane (23). We hypothesize that the higher Zn content in the upper conducting airway epithelium reflects the greater need to protect these cells from foreign inhaled particles and other noxious agents.
It is important to mention that Zinquin binds to intracellular labile
Zn with a dissociation constant (Kd) of
107 M. By labile, we mean the pools normally
1) participating in Zn fluxes and ionic exchange,
2) most readily altered in imbalances of Zn homeostasis, and
3) not tightly bound to metalloproteins (Kd
10
11 to 10
13
M) (35). The distribution of total Zn is not known but is
likely to be similar to the distribution of metalloenzymes in these
cells and, therefore, is uniformly distributed throughout the cellular organelles (32). The depletion of intracellular Zn was
achieved with TPEN, a membrane-permeable heavy metal chelator that
binds intracellular labile Zn [log association constant
(Ka) 15.6] via appropriately spaced nitrogens
in the ring structure, but has minimal affinity for calcium or
magnesium (4). This chelator has a much higher affinity
for Zn than does Zinquin (log Ka1 6.4 and log
Ka2 7.1, where Ka1 and
Ka2 are the association constants for the
formation of Zinquin-Zn and (Zinquin)2-Zn complexes,
respectively) and quenches Zinquin fluorescence in solution
and within cells (35).
We now need to determine and compare the Zn distribution in respiratory tissues of humans and other species. Our studies have shown that upper respiratory epithelial cells are rich in Zn, leading us to speculate that constant insults to this tissue by proteases and cytokines from inflammatory cells would cause a significant loss of Zn. This may contribute to the known hypozincemia that is a reported feature of asthma (10, 11, 17). In a similar way, Zn deficiency also arises in burn patients where there is enhanced cell turnover in the damaged skin (28).
As discussed earlier (14), Zn may have a special antioxidant role in the vicinity of mitochondria, which release potentially damaging oxyradicals during the generation of energy. This is a particular hazard for the ciliated cells lining the airways because these cells rely heavily on this energy for rapid beating of the cilia. Zn may exert a protective, antioxidant role in these cells against oxidants both inhaled down the airways and released from mitochondria as a by-product of oxidative metabolism. It will be interesting to determine whether other antioxidants (e.g., glutathione and Cu/Zn superoxide dismutase) are similarly concentrated in this apical region.
Apical Zn may be derived from Zn reservoirs existing in a possible vesicular and perinuclear pattern. This was particularly evident in Zn-supplemented transformed cells when viewed at high magnification. In the primary epithelial cells, the perinuclear concentration of Zn was most pronounced on the apical side of the nucleus and is reminiscent of the distribution of Zn in secretory cells such as the Zn-rich pancreatic islets (37). Whether any of the apical Zn in the ciliated cells is contained within secretory granules and is destined for secretion into the pericellular fluid is not clear. Measurement of the Zn content in this fluid should be informative. The confocal studies also indicated a significant content of labile Zn within the cilia themselves. Zn is a well-known stabilizer of microtubules (26) and may have a similar role in the axonemes of the cilia.
We propose that the major function of apical labile Zn in the ciliated cells is cytoprotective and involves suppression of cell death by apoptosis and perhaps also necrosis. It is interesting to note that our observed distribution of labile Zn in these cells closely matches that of the major executioner enzyme in apoptosis, caspase-3, as detected by immunocytochemistry in human tracheobronchial epithelium (18). The close proximity of labile Zn, with procaspase-3 and the rapid activation of this enzyme after Zn depletion, raises the question of whether apical cytoplasmic Zn can act to suppress the induction of this caspase. A similar conclusion was reached by Aiuchi et al. (2) and Wolf and Eastman (34) in nonrespiratory cell lines. Zn is only a weak suppressor of active caspase-3 but a potent suppressor of caspase-6 at low micromolar concentrations (29) and possibly also of caspase-9 (34). However, both of these caspases contribute to the activation of caspase-3 (29, 30, 34). Caspase-6 was activated in our Zn-depleted respiratory epithelial cells, but there was no evidence that this event occurred before the activation of caspase-3. One implication of the relatively late (after 3 h) increase in caspase-6 activity is that this enzyme is also required during the effector phase where it cleaves lamin proteins, leading to nuclear collapse (30). Because it has been shown that caspase-3 and caspase-6 act synergistically to trigger morphological changes in apoptosis (30), the induction of both caspases in Zn-depleted cells is consistent with other models of apoptosis. Activated caspase-9, on the other hand, was undetected in our Zn-deprived cells. This could be due to an absence of this particular enzyme in primary sheep ciliated epithelial cells. These issues are currently being pursued with cell-free extracts.
Induction of high rates of apoptosis has previously been observed in the gastrointestinal epithelium of Zn-deficient animals (12). Furthermore, Zn deficiency rendered this tissue more susceptible to damage by toxins (9). Similar in vivo animal studies are required to answer the question of whether the respiratory epithelium is also affected in Zn deficiency. Our experiments in vitro with primary and malignant respiratory epithelial cells support the hypothesis that Zn deficiency renders the airway epithelium more susceptible to damage by apoptosis-inducing toxins. That this occurred with two different types of an apoptosis inducer is consistent with a generalized effect of Zn depletion on increased susceptibility to apoptosis. It is important to remember that once caspase-3 has been activated in a damaged cell, then the cell is committed to apoptosis regardless of the presence of Zn. However, Zn will suppress the death of those cells that have not yet had their caspase-3 enzymes activated.
Our findings suggest that Zn suppresses caspase activation in these cells and might have potential therapeutic implications in epithelial protection. Studies are now required to establish the intracellular labile Zn status of airway epithelial cells in vivo in healthy and asthmatic subjects and to correlate these levels with altered caspase activity and susceptibility to apoptosis. In conclusion, we have shown an unexpectedly high concentration of labile Zn lining the airway epithelium in pigs and sheep and propose that this Zn exerts a largely protective role against cell death induced by oxidants derived either from the lumen or as by-products of oxidative metabolism.
![]() |
ACKNOWLEDGEMENTS |
---|
We are grateful to Dr. Darryl Knight and Dr. Prue Cowled for helpful discussions. Many thanks to Dr. Peter Kolesik for assistance with the ultraviolet laser confocal microscope, John Brealey for electron microscopy studies, Ken Porter for animal tissues, and Suzie Dosljak (Department of Immunohistochemistry) for assistance with the cryostat sections.
![]() |
FOOTNOTES |
---|
We acknowledge the Queen Elizabeth Hospital Research Foundation for financial support.
A. Q. Truong-Tran was the recipient of a Benjamin Poulton University of Adelaide Postgraduate Scholarship.
Address for reprint requests and other correspondence: P. D. Zalewski, Dept. of Medicine, Univ. of Adelaide, The Queen Elizabeth Hospital, Woodville, South Australia 5011, Australia (E-mail: peter.zalewski{at}adelaide.edu.au).
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.
Received 17 February 2000; accepted in final form 13 June 2000.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Agren, M,
Chvapil M,
and
Franzen L.
Enhancement of re-epithelialization with topical zinc oxide in porcine partial-thickness wounds.
J Surg Res
50:
101-105,
1991[ISI][Medline].
2.
Aiuchi, T,
Mihara S,
Nakaya M,
Masuda Y,
Nakajo S,
and
Nakaya K.
Zinc ions prevent processing of caspase-3 during apoptosis induced by geranylgeraniol in HL-60 cells.
J Biochem (Tokyo)
124:
300-303,
1998[Abstract].
3.
Antoshina, E,
and
Ostrowski LE.
TGF1 induces growth arrest and apoptosis but not ciliated differentiation in rat tracheal epithelial cell cultures.
In Vitro Cell Dev Biol Anim
33:
212-217,
1997[Medline].
4.
Arslan, P,
Di Virgilio F,
Beltrame M,
Tsien RY,
and
Pozzan T.
Cytosolic Ca2+ homeostasis in Ehrlich and Yoshida carcinomas.
J Biol Chem
260:
2719-2727,
1985[Abstract].
5.
Carney, DN,
Gazdar AF,
Bepler G,
Guccion JG,
Marangos PJ,
Moody TW,
Zweig MH,
and
Minna JD.
Establishment and identification of small cell lung cancer cell lines having classic and variant features.
Cancer Res
45:
2913-2923,
1985[Abstract].
6.
Coyle, P,
Zalewski PD,
Philcox JC,
Forbes IJ,
Ward AD,
Lincoln SF,
Mahadevan I,
and
Rofe AM.
Measurement of zinc in hepatocytes using a fluorescent probe, Zinquin: relationship to metallothionein and intracellular zinc.
Biochem J
303:
781-786,
1994[ISI][Medline].
7.
Da Silva, JJ,
and
Williams RJ.
Zinc: Lewis acid catalysis and regulation.
In: The Biological Chemistry of the Elements, edited by Williams RJ.. Oxford, UK: Clarendon, 1991, p. 299-318.
8.
De Raeve, HR,
Thunnissen FB,
Kaneko FT,
Guo FH,
Lewis M,
Kavuru MS,
Secic M,
Thomassen MJ,
and
Erzurum SC.
Decreased Cu,Zn-SOD activity in asthmatic airway epithelium: correction by inhaled corticosteroid in vivo.
Am J Physiol Lung Cell Mol Physiol
272:
L148-L154,
1997
9.
Dinsdale, D,
and
Williams RB.
The enhancement by dietary zinc deficiency of the susceptibility of the rat duodenum to colchicine.
Br J Nutr
37:
135-142,
1977[ISI][Medline].
10.
Di Toro, R,
Galdo Capotorti G,
Gialanella G,
Miraglia del Giudice M,
Moro R,
and
Perrone L.
Zinc and copper status of allergic children.
Acta Paediatr Scand
76:
612-617,
1987[ISI][Medline].
11.
El-Kholy, MS,
Gas Allah MA,
el-Shimi S,
el-Baz F,
el-Tayeb H,
and
Abdel-Hamid MS.
Zinc and copper status in children with bronchial asthma and atopic dermatitis.
J Egypt Public Health Assoc
65:
657-668,
1990[Medline].
12.
Elmes, ME.
Apoptosis in the small intestine of zinc-deficient and fasted rats.
J Pathol
123:
219-224,
1977[ISI][Medline].
13.
Hagimoto, N,
Kuwano K,
Miyazaki H,
Kunitake R,
Fujita M,
Kaneko Y,
and
Hara N.
Induction of apoptosis and pulmonary fibrosis in mice in response to ligation of Fas antigen.
Am J Respir Cell Mol Biol
17:
272-278,
1997
14.
Halliwell, B,
Gutteridge JM,
and
Cross CE.
Free radicals, antioxidants, and human disease: where are we now?
J Lab Clin Med
119:
598-620,
1992[ISI][Medline].
15.
Harding, AJ,
Dreosti IE,
and
Tulsi RS.
Zinc deficiency in the 11 day rat embryo: a scanning and transmission electron microscope study.
Life Sci
42:
889-896,
1988[ISI][Medline].
16.
Hennig, B,
Meerarani P,
Ramadass P,
Toborek M,
Malecki A,
Slim R,
and
McClain CJ.
Zinc nutrition and apoptosis of vascular endothelial cells: implications in atherosclerosis.
Nutrition
15:
744-748,
1999[ISI][Medline].
17.
Kadrabova, J,
Mad'aric A,
Podivinsky F,
Gazdik F,
and
Ginter F.
Plasma zinc, copper and copper/zinc ratio in intrinsic asthma.
J Trace Elem Med Biol
10:
50-53,
1996[ISI][Medline].
18.
Krajewska, M,
Wang HG,
Krajewski S,
Zapata JM,
Shabaik A,
Gascoyne R,
and
Reed JC.
Immunohistochemical analysis of in vivo patterns of expression of CPP32 (caspase-3), a cell death protease.
Cancer Res
57:
1605-1613,
1997[Abstract].
19.
Laitinen, LA,
Heino M,
Laitinen A,
Kava T,
and
Haahtela T.
Damage of the airway epithelium and bronchial reactivity in patients with asthma.
Am Rev Respir Dis
131:
599-606,
1985[ISI][Medline].
20.
Lieber, M,
Smith B,
Szakal A,
Nelson-Rees W,
and
Todaro G.
A continuous tumor-cell line from a human lung carcinoma with properties of type II alveolar epithelial cells.
Int J Cancer
17:
62-70,
1976[ISI][Medline].
21.
Mahadevan, I,
Kimber MC,
Lincoln SF,
Tiekink ER,
Ward AD,
Betts WH,
Forbes IJ,
and
Zalewski PD.
The synthesis of Zinquin ester and Zinquin acid, Zn (II)-specific fluorescing agents for use in the study of biological Zn (II).
Aust J Chem
49:
561-568,
1996[ISI].
22.
Marini, M,
Vittori E,
Hollemborg J,
and
Mattoli S.
Expression of the potent inflammatory cytokines, granulocyte-macrophage-colony-stimulating factor and interleukin-6 and interleukin-8, in bronchial epithelial cells of patients with asthma.
J Allergy Clin Immunol
89:
1001-1009,
1992[ISI][Medline].
23.
McMahon, RJ,
and
Cousins RJ.
Mammalian zinc transporters.
J Nutr
128:
667-670,
1998
24.
Medina, V,
Edmonds B,
Young GP,
James R,
Appleton S,
and
Zalewski PD.
Induction of caspase-3 protease activity and apoptosis by butyrate and trichostatin A (inhibitors of histone deacetylase): dependence on protein synthesis and synergy with a mitochondrial/cytochrome c-dependent pathway.
Cancer Res
57:
3697-3707,
1997[Abstract].
25.
Nakajima, Y,
Aoshiba K,
Yasui S,
and
Nagai A.
H2O2 induces apoptosis in bovine tracheal epithelial cells in vitro.
Life Sci
64:
2489-2496,
1999[ISI][Medline].
26.
Oteiza, PI,
Hurley LS,
Lonnerdal B,
and
Keen CL.
Effects of marginal zinc deficiency on microtubule polymerization in the developing rat brain.
Biol Trace Elem Res
24:
13-23,
1990[ISI][Medline].
27.
Polunovsky, VA,
Chen B,
Henke C,
Snover D,
Wendt C,
Ingbar DH,
and
Bitterman PB.
Role of mesenchymal cell death in lung remodeling after injury.
J Clin Invest
92:
388-397,
1993[ISI][Medline].
28.
Solomons, NW.
Zinc and copper.
In: Modern Nutrition in Health and Disease (7th ed.), edited by Shils ME,
and Young VR.. Philadelphia, PA: Lea & Febiger, 1988, p. 238-262.
29.
Stennicke, HR,
and
Salvesen GS.
Biochemical characteristics of caspases-3, -6, -7, and -8.
J Biol Chem
272:
25719-25723,
1997
30.
Takahashi, A,
Alnemri ES,
Lazebnik YA,
Fernandes-Alnemri T,
Litwack G,
Moir RD,
Goldman RD,
Poirier GG,
Kaufmann SH,
and
Earnshaw WC.
Cleavage of lamin A by Mch2 alpha but not CPP32: multiple interleukin 1 beta-converting enzyme-related proteases with distinct substrate recognition properties are active in apoptosis.
Proc Natl Acad Sci USA
93:
8395-8400,
1996
31.
Takizawa, H,
Romberger D,
Beckmann JD,
Matsuda T,
Eccleston-Joyner C,
Shoji S,
Rickard KA,
Claassen LR,
Ertl RF,
Linder J,
and
Rennard SI.
Separation of bovine bronchial epithelial cell subpopulations by density centrifugation: a method to isolate ciliated and nonciliated cell fractions.
Am J Respir Cell Mol Biol
3:
553-562,
1990[ISI][Medline].
32.
Vallee, BL,
and
Falchuk KH.
The biochemical basis of zinc physiology.
Physiol Rev
73:
79-118,
1993
33.
Wen, LP,
Madani K,
Fahrni JA,
Duncan SR,
and
Rosen GD.
Dexamethasone inhibits lung epithelial cell apoptosis induced by IFN- and Fas.
Am J Physiol Lung Cell Mol Physiol
273:
L921-L929,
1997
34.
Wolf, CM,
and
Eastman A.
The temporal relationship between protein phosphatase, mitochondrial cytochrome c release, and caspase activation in apoptosis.
Exp Cell Res
247:
505-513,
1999[ISI][Medline].
35.
Zalewski, PD,
Forbes IJ,
and
Betts WH.
Correlation of apoptosis with change in intracellular labile zinc, using Zinquin, a new specific fluorescent probe for zinc.
Biochem J
296:
403-408,
1993[ISI][Medline].
36.
Zalewski, PD,
Jian X,
Soon LL,
Breed WG,
Seamark RF,
Lincoln SF,
Ward AD,
and
Sun FZ.
Changes in distribution of labile zinc in mouse spermatozoa during maturation in the epididymis assessed by the fluorophore Zinquin.
Reprod Fertil Dev
8:
1097-1105,
1996[ISI][Medline].
37.
Zalewski, PD,
Millard SH,
Forbes IJ,
Kapaniris O,
Slavotinek A,
Betts WH,
Ward AD,
Lincoln SF,
and
Mahadevan I.
Video image analysis of labile zinc in viable pancreatic islet cells using a specific fluorescent probe for zinc.
J Histochem Cytochem
42:
877-884,
1994