1 Department of Pathology, University of Vermont, Burlington, Vermont 05405; and 2 Biophysics Research Institute, Medical College of Wisconsin, Milwaukee, Wisconsin 53226
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ABSTRACT |
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Nitric oxide (NO ·) is important in the regulation of mitochondrial function, cell signaling, and gene expression. To elucidate how endogenous NO · regulates the function of airway epithelial cells, we used carboxy-PTIO, a hydrophilic, negatively charged NO · trap, to scavenge NO · from rat lung epithelial (RLE) and rat pleural mesothelial (RPM) cells and to determine the elicitation of cell cycle alterations, apoptosis, and oxidative stress. The reaction of NO · with PTIO causes the formation of PTI, which is measured by electron spin resonance (ESR) and is a quantitative measure of NO · formation. ESR spectroscopy revealed the production of NO · in RLE or RPM cells over a period from 1 to 24 h of exposure, indicating scavenging of NO · by PTIO. Cycle analyses in confluent RLE or RPM cells revealed two- to threefold increases in S and G2/M phases after exposure to 100-200 µM PTIO as well as increases in the fraction of cells undergoing apoptosis. Direct addition of PTI to cells failed to elicit cell cycle perturbations or apoptosis. The guanylyl cyclase inhibitor ODQ mimicked the effects of PTIO. 8-Bromo-cGMP but not 8-bromo-cAMP ameliorated the PTIO- or ODQ-mediated cell cycle perturbations and apoptosis, suggesting that cGMP-dependent pathways are involved in these cell cycle perturbations. Treatment of log-phase cells with PTIO resulted in more dramatic cell cycle perturbations compared with cells treated at confluence. Assessment of 5-bromo-2'-deoxyuridine incorporation to measure DNA synthesis demonstrated decreases in PTIO-treated compared with sham cells in addition to a cell cycle arrest in late S or G2/M phase. Last, incubation with dichlorofluorescin diacetate revealed oxidative stress in PTIO- but not in PTI-exposed RLE or RPM cells. We conclude that the depletion of endogenous NO · induces oxidative stress, cell cycle perturbations, and apoptosis. Our findings illustrate the importance of endogenous NO · in the control of cell cycle progression and survival of pulmonary and pleural cells and that a critical balance between NO · and superoxide may be necessary for these physiological events.
nitronyl nitroxide; epithelial cell; mesothelial cell
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INTRODUCTION |
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NITROGEN MONOXIDE [nitric oxide (NO ·)] is a free radical gas with critical physiological and homeostatic functions. In the lung, NO · plays an important role in the control of vascular and bronchial tone and neuroendocrine regulation of airway mediator release (16, 45). NO · can be measured in human airways, and its levels are increased in patients with pulmonary diseases including asthma (5). NO · is a potent smooth muscle relaxant that results in dilation of bronchioles and blood vessels. Because of these properties, NO · is used clinically to treat pulmonary hypertension (9).
NO ·, formed in large quantities during inflammation, is
thought to contribute to tissue damage because it has the potential to
react with iron sulfur clusters in proteins to abrogate their function
(17, 41). Modification of aconitase, cytochrome
c oxidase, and ribonucleotide
reductase occurs after NO · exposure (17, 41). Addition of
NO · donors to a range of cell types, including pulmonary
epithelial cells, results in apoptosis (23), illustrating the toxic
properties of NO ·. It is likely that the kinetics of
NO · released by exogenously added NO · donors as well as the steady-state concentrations of NO · achieved in
these models are often not physiological and may confound our
understanding of the physiological mechanism of action of
NO · in cells. This is further complicated by superoxide
(O2·), a ubiquitous reaction
product of phagocytic cells during inflammation and a minor product of
mitochondrial oxidative metabolism (1%) (37). NO · and
O
2· react rapidly (second-order rate constant = 6.7 × 109
M/s) to generate peroxynitrite
(ONOO
), which can account
for some of the toxic effects associated with NO · (7). In
contrast to the cytotoxic effects of NO ·, recent studies
(10, 20, 32, 35, 38, 40) documented a protective role of endogenous
NO · against apoptosis and
O
2·-mediated cell death.
Pharmacological inhibition of nitric oxide synthases (NOSs) leads to
enhanced O
2· and
H2O2
production (18, 34) and promotes adherence of inflammatory cells to
endothelium (34). These data suggest that a critical balance of
NO · and O
2· is required
for normal cell functioning and prevention of inflammation.
Airway and alveolar epithelial, neuroendocrine, smooth muscle, and pleural mesothelial cells have both constitutive and inducible forms of NOS and can be activated to release NO · on exposure to a wide array of stimuli (3, 16). The function of endogenous NO · in these cell types is not clear. It is currently believed that a critical balance of reactive oxygen species or reactive nitrogen species exists in cells that is important for the redox regulation and control of cell signaling, activation of transcription factors, and gene expression (11, 42). For instance, it has been demonstrated that NO · activates Ras (27) and mitogen-activated protein kinases (28) that are critical in the control of the transcription factor activator protein-1, which may regulate apoptosis, proliferation, or survival (2). Endogenous NO · may therefore be important in the control of activities of these regulatory proteins. Furthermore, numerous biological activities of NO · are mediated as a result of activation of guanylate cyclase (29). Control of cGMP-dependent pathways by NO · may therefore be important in the regulation of normal cell physiology.
To date, few methods are available for the sensitive quantitation of NO · in cellular and biological systems (25). For instance, one study (36) involved the use of pharmacological inhibitors of NOSs, which may have side effects and display different sensitivities toward inhibition of the distinct NOS isoform (36). To circumvent these potential problems, we focused on nitronyl nitroxides, which are known scavengers of NO · (1, 36). In this study, we used the hydrophilic, negatively charged nitronyl nitroxide 1H-imidazol-1-yloxy,2-(4-carboxyphenyl)-4,5-dihydro-4,4,5,5-tetramethyl-,3-oxide (carboxy-PTIO) as a trap for NO · and investigated the functional consequences of NO · depletion in pulmonary cells. This compound has been used in a biophysical study (25) to quantitate NO · formation in vitro. The objectives of the present study were to evaluate how NO · regulates the function of rat lung alveolar type II epithelial (RLE) and rat pleural mesothelial (RPM) cells with PTIO to deplete NO · from cells. Our results with PTIO demonstrate the formation of NO · in pulmonary and pleural cells and that depletion of endogenous NO · leads to cell cycle perturbations, apoptosis, and oxidative stress. The effects may be due to alterations of guanylyl cyclase and cGMP because inhibition of guanylyl cyclase mimicked the effects of PTIO, whereas 8-bromo-cGMP (8-BrcGMP) but not 8-bromo-cAMP (8-BrcAMP) ameliorated the apoptotic response associated with NO · depletion. Our data also demonstrate that a critical level of endogenous NO · is required for normal cell cycle progression, survival, and prevention of oxidative stress.
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EXPERIMENTAL PROCEDURES |
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Cell culture and exposure to test agents. A line of spontaneously transformed RLE cells was kindly provided by Dr. Kevin Driscoll (Procter and Gamble, Cincinnati, OH) and has been characterized elsewhere (13). RLE cells were maintained in DMEM-Ham's F-12 medium supplemented with 7% newborn bovine serum. Primary isolates of RPM cells were obtained by gentle scraping of the parietal pleura of Fischer 344 rats (24). RPM cells were propagated in DMEM-Ham's F-12 medium containing 10% fetal bovine serum supplemented with hydrocortisone, insulin, transferrin, and selenium. Cells were grown to confluency, and the medium was switched to 1% serum for 24 h before addition of the test agents. In selected experiments, RLE cells were exposed to PTIO during log-phase growth. Carboxy-PTIO was obtained from Cayman Chemical (Ann Arbor, MI). PTI was synthesized by the reaction of PTIO with sodium nitrite and purified by column chromatography (43). PTI does not react with NO · as reported earlier (21). PTIO or PTI was dissolved in 50 mM NaOH immediately before addition to the cell cultures and was used at concentrations of 50-200 µM. The selective NO ·-sensitive inhibitor of guanylyl cyclase 1H-[1,2,4]oxadiazole[4,3-a]quinoxalin-1-one (ODQ) (15,33) was purchased from Alexis Biochemicals (San Diego, CA), and 8-BrcGMP and 8-BrcAMP were from Calbiochem (La Jolla, CA).
Trapping of NO · with PTIO. After 2, 4, 8, or 24 h of exposure to PTIO or PTI, 100 µl of the cell culture medium were removed and quickly frozen in liquid nitrogen for electron spin resonance (ESR) analysis (1, 25). The cells were trypsinized briefly, centrifuged for 10 min at 1,500 rpm at 4°C, washed once in PBS, and frozen in liquid nitrogen for ESR analysis. ESR spectra were recorded at room temperature on a Varian E-109 spectrometer operating at 9.5 gZH and with a 100-kHz field modulation equipped with a TE102 cavity. Samples were taken in 100-µl capillaries and inserted into 4-mm quartz tubes for ESR analysis. Reoxidation of hydroxylamine to nitroxide was achieved by adding ferricyanide (1 µl of a 100 mM stock solution) to 100 µl of the incubation mixture.
Flow cytometry for the determination of cell cycle perturbations and apoptosis. Twenty-four hours after the addition of PTIO or PTI, the cells were trypsinized briefly, centrifuged at 1,500 rpm for 10 min, and resuspended in a propidium iodide-staining solution (3.75 mM sodium citrate, 0.1% Triton X-100, 30 µg/ml of RNase A, and 20 µg/ml of propidium iodide) for flow cytometric analysis of the DNA content of 10,000 cells. This procedure determines the proportion of cells in the G0/G1, S, and G2/M phases of the cell cycle. In addition, the apoptotic fraction is recognized by the patterns of forward and side scatter and the sub-G0/G1 DNA content that results from cleavage of DNA during apoptosis (12, 23). Induction of apoptosis was further confirmed by staining the cells with annexin V, which evaluates the presence of phosphatidylserine residues on the outer membrane as a result of the loss of cell membrane phospholipid asymmetry during the apoptotic process. Flow cytometric analysis of annexin V-phycoerythrin-positive cells that are excluding propidium iodide was performed according to manufacturer's instructions (R&D Systems, Minneapolis, MN). Last, apoptosis was confirmed by in situ techniques and staining the DNA with 4,6-diamidino-2-phenylindole (DAPI) according to procedures described elsewhere (19).
Incorporation of 5-bromo-2'-deoxyuridine to assess DNA synthesis. In selected experiments, cells were incubated with 10 µM 5-bromo-2'-deoxyuridine (BrdU) for 16 h before they were harvested. The cells were obtained by brief trypsinization, and 0.3 × 106 cells were utilized for BrdU-propidium iodide-staining experiments. The cells were washed and resuspended in 150 µl of PBS containing 1% bovine serum albumin and incubated for 15 min at 4°C on a rotating platform after the addition of 300 µl of lysis propidium iodide solution (0.1% Triton X-100, 20 µg/ml of propidium iodide, 0.2 mg/ml of RNase A, and 0.5 mM EDTA). Subsequently, 75 µl of 1 N HCl were added for 30 min at room temperature, followed by the addition of 225 µl of 1 N Tris base for 5 min. One microliter of fluorescein-conjugated anti-BrdU antibody (Boehringer Mannheim, Indianapolis, IN) was added for 15 min at 4°C on a rotating platform and samples were analyzed by flow cytometry.
Assessment of oxidative stress. After various times during the exposure to PTIO or PTI, RLE or RPM cells were harvested by brief trypsinization as described above. Cells were subsequently incubated with 10 µM carboxydichlorodihydrofluorescein diacetate [dichlorofluorescin diacetate (DCF); Molecular Probes, Eugene, OR] for 30 min at 37°C, and 10,000 cells were analyzed by flow cytometry to determine the oxidation of DCF that occurs during oxidative stress (46).
Statistical analyses. The results were analyzed with ANOVA with the Student-Newman-Keuls procedure to adjust for multiple comparisons. Experiments were repeated two times, and the data are expressed as means ± SE.
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RESULTS |
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We first monitored the production of NO · by RLE cells after incubation with 100 µM PTIO. Figure 1A shows the ESR spectra of the cell culture medium taken at different time intervals. The spectrum of PTIO consists of a five-line signal with an intensity ratio of 1:2:3:2:1. On the basis of published data (1, 25), this spectrum is attributed to an electron interacting with two equivalent nitrogens (a1N = 8.2 G; a2N = 8.2 G; where a is a hyperfine coupling constant). With a prolonged incubation period, the spectral intensity of PTIO gradually decreased, and the ESR spectrum disappeared completely after a 24-h incubation period. The loss in signal intensity is attributed to the reduction in PTIO and PTI to the corresponding hydroxylamine that occurs in cells. This can be overcome by the addition of ferricyanide, a commonly used oxidizing agent that converts hydroxylamines to nitroxides. Figure 1B shows the ESR spectra of samples treated with ferricyanide. The line positions due to PTI are marked. The PTI signal that results from the reaction of NO · with PTIO increased with time and reached a maximum after 24 h of exposure (Fig. 1C). The ESR signal obtained after 8 and 24 h is a composite of both PTIO and PTI. When the cell pellets were evaluated, no detectable ESR spectrum of PTIO or PTI was obtained (data not shown). Pretreatment of cells with N-nitro-L-arginine methyl ester (L-NAME; 5 mM) inhibited formation of PTI by ~50%, suggesting the involvement of NO · in the conversion of PTIO to PTI (data not shown). These findings demonstrate that RLE cells produce NO · and that this can be measured sensitively by incubation of the cells with PTIO. Furthermore, trapping of NO · from RLE cells by PTIO was maximal after 8-24 h of exposure (Fig. 1C).
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To determine whether trapping of NO · from RLE cells perturbs the cell cycle, we first examined alterations in cell cycle distributions and apoptosis by flow cytometry. The results in Fig. 2A illustrate the increases in the S and G2/M phases of the cell cycle (fractions C and D) of RLE cells exposed to 100 µM PTIO for 24 h. Furthermore, a hypodiploid fraction characteristic of apoptosis (Fig. 2A, fraction A) is apparent in PTIO-exposed cells. None of these changes are apparent when the cells were incubated with PTI, which cannot trap NO · from cells. Quantitation of these findings (Fig. 2B) revealed three- to eightfold increases in RLE cells undergoing apoptosis and two- to threefold increases in the fraction of cells in the S or G2/M phase after exposure to 50-200 µM PTIO.
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To determine whether the PTIO-induced responses were specific to lung epithelial cells, we also evaluated cell cycle alterations in RPM cells. As illustrated in Fig. 2C, exposure of RPM cells to PTIO increased the fraction of cells undergoing apoptosis and in the S and G2/M phases of the cell cycle. A less striking but significant increase in the apoptotic fraction was observed after exposure of RPM cells to PTI but was not confirmed by other criteria of apoptosis. To further validate the induction of apoptosis by PTIO, we evaluated the occurrence of phosphatidylserine residues on the outer surface of RLE cells by staining the cells with annexin V and found that 0.4% of untreated control cells were annexin V positive. Annexin V-positive RLE cells increased to 3.1 and 7.6% in cultures treated with 100 and 200 µM PTIO, respectively, for 24 h, similar to trends observed with propidium iodide staining. Quantitation of apoptotic cells with image analysis and the DNA stain DAPI (19) revealed 2.5-fold increases in RLE cells exposed to 100 µM PTIO (P < 0.05 by ANOVA), whereas PTI failed to increase apoptotic cells, identical to findings described in Fig. 2B. Consistent with findings that PTIO increases the proportion of RLE cells in the S and G2/M phases of the cell cycle, we observed numerous cells exhibiting mitotic spindle formation, characteristic of the M phase of the cell cycle (data not shown).
Numerous effects of NO · are attributed to the NO ·-mediated activation of guanylyl cyclase and increases in cGMP levels, which activate downstream cascades. To determine whether guanylyl cyclase is involved in the cell cycle perturbations and apoptosis, we incubated cells with the specific inhibitor of NO ·-sensitive guanylyl cyclase ODQ (15, 33). Figure 3A illustrates that exposure of RLE cells to 50 µM ODQ resulted in identical perturbations of the cell cycle and apoptosis that were observed after PTIO exposure. As expected, 1 mM 8-BrcGMP almost completely reversed the ODQ-mediated cell cycle perturbations, whereas 1 mM 8-BrcAMP had no effect. We next determined whether the addition of 8-BrcGMP could alleviate the PTIO-induced effects in RLE cells. As is shown in Fig. 3B, exposure of the cells to 8-BrcGMP but not to 8-BrcAMP ameliorated the PTIO-induced apoptosis. However, 8-BrcGMP did not completely abolish the response to PTIO, suggesting that both cGMP-dependent and -independent pathways may be associated with the cell cycle perturbations observed after NO · depletion.
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The cell cycle perturbations by PTIO and ODQ reported above are observed in confluent cultures of pulmonary and pleural cells. We therefore hypothesized that the effects of NO · depletion would be enhanced in actively growing cultures of epithelial cells. We next exposed log-phase RLE cultures to PTIO and determined the occurrence of cell cycle perturbations. As illustrated in Fig. 4A, 100 µM PTIO caused increases in the S (fraction J) and G2/M (fraction K) phases after 24 h of exposure that are still apparent after 48 h. Although a sub-G0/G1 population (Fig. 4A, fraction H) is not readily apparent after 24 h of exposure to PTIO, after 48 h of exposure, striking increases in the sub-G0/G1 phase (fraction H) occur, indicative of apoptosis. Figure 4B demonstrates that the effects of PTIO in subconfluent cultures are partially reversible by 500 µM 8-BrcGMP but not by 8-BrcAMP as was apparent in confluent cultures.
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The increases in the S and G2/M fractions may be indicative of a block in cell cycle progression. To address this, we used the thymidine analog BrdU to assess DNA synthesis after treatment with PTIO. As shown in Fig. 4C, exposure to PTIO causes an ~50% decrease in DNA synthesis, accompanied by similar decreases in total cell number (data not shown). The PTIO-induced inhibition of DNA synthesis was almost completely reversed by 8-BrcGMP but not by 8-BrcAMP. Although 8-BrcAMP caused a 25% decrease in BrdU incorporation, the magnitude of PTIO-mediated decreases was similar to that of nonpreexposed cells. We next performed BrdU flow cytometric experiments combined with propidium iodide staining to identify the cell cycle arrest point in response to PTIO. As demonstrated in Fig. 4D, most sham-treated cells have traversed the cell cycle and are BrdU positive throughout the cell cycle. In contrast, after 24 h of PTIO exposure, a large fraction of BrdU-positive cells accumulates in the late S and, more predominantly, G2/M phases. Evaluation at later time points (48 and 72 h) also demonstrates this accumulation in the late S and G2/M phases. These results indicate that the PTIO-induced cell cycle arrest occurs late in the S phase or at the G2/M boundary.
Depletion of endogenous NO · may disrupt the balance between
NO · and O2· and may
promote O
2·dependent oxidative stress. We therefore determined whether oxidation of DCF was
enhanced in response to PTIO, which reflects the increased production
of oxidants, including
H2O2,
in the cells (46). The flow cytometric data illustrate the occurrence
of oxidative stress in RLE (Fig.
5A) and
RPM cells (Fig. 5B) exposed to PTIO.
Exposure of cells to PTI did not affect DCF oxidation in either cell
type.
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DISCUSSION |
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Pulmonary epithelial and mesothelial cells release NO · and
other oxidants in response to a wide variety of agents such as cytokines, inflammatory particulates, or inhaled microorganisms (3,
45). Release of these oxidants is critical in initial protection but
may aggravate disease when a prolonged release occurs. Although
numerous studies (3, 45), reported an increased release
of NO · from cells exposed to a multitude of stimuli, information on the function of endogenous NO · in pulmonary
epithelial or pleural mesothelial cells is sparse. In the present
study, we evaluated whether NO · is produced endogenously in
unexposed pulmonary epithelial and mesothelial cells. We also
determined the functional ramifications of trapping of physiological
levels of NO · from pulmonary epithelial or pleural
mesothelial cells. Direct detection of NO · in cellular
systems is difficult. Here we used the nitronyl nitroxide PTIO to trap
NO · from cells. PTIO has been used extensively in
biophysical systems and selectively reacts with NO ·. This
reaction yields the formation of PTI, which can be measured by ESR (1,
25). The use of a nitronyl nitroxide has the advantage over other
approaches in that it reacts very specifically with NO · and
allows detection of NO · in very low concentrations, which is
not possible with the use of other reagents including the Griess
reaction. To our knowledge, no other oxidants (O2·,
NO2 ·, or
· OH) react with PTIO to form PTI. Based on the partial
inhibitory effect observed in the presence of
L-NAME, we attribute the
transformation of PTIO to PTI to the trapping of NO ·,
although we cannot rule out cellular metabolism of PTIO to PTI by an as
of yet undiscovered mechanism. Our approach also differs from other
reports that used pharmacological inhibitors of NOS to downregulate
production of NO · but may have side effects and inhibit NOSs
differentially (36). Exposure of RLE or RPM cells to PTIO resulted in
the formation of PTI, the product of the reaction of PTIO with
NO ·, indicating that pulmonary cells produce detectable
levels of NO ·.
Trapping of NO · had multiple functional consequences that
include cell cycle perturbations, decreases in DNA synthesis,
apoptosis, and enhanced oxidative stress. The disturbances of the
balance of endogenous NO · and
O2· in pulmonary cells may affect
signaling cascades and elicit these phenotypic consequences. Our
findings agree with a previous report (30) that showed that inhibition
of NOS by pharmacological agents sensitizes endothelial cells to
undergo apoptosis.
At present, the exact source of endogenous
O2· formation is unknown. It is
likely that the mitochondria are involved because these organelles are
a major site for formation of reactive oxygen species and reactive
nitrogen species in nonimmune cells. A recent study (6) illustrated
that an NOS is present in the mitochondria where it may control levels
of O
2· formation via the
production of ONOO
.
Increased O
2· production in
mitochondria as a result of the trapping of NO · may induce
the mitochondrial transition pore and cause release of cytochrome
c into the cytoplasm, events that are
critical to the initiation of apoptosis (26). Our findings suggest an
important role for endogenous NO · in the control of the
apoptotic response.
The alterations in cell cycle distribution observed after exposure to an NO · trap are intriguing. Enhanced H2O2 formation may be responsible for some of these effects. A previous report (42) demonstrated that H2O2 can enhance cell proliferation and thus could stimulate cells to enter the S phase. However, in the present study, we observed decreases in DNA synthesis and cell number in response to NO · depletion, in contrast to proliferative responses that may be expected after exposure to low levels of peroxides (42). Although the reaction between PTIO and NO · produces the NO2 · radical, which may contribute to oxidative stress, it is likely that NO2 · is scavenged by the reduced form of PTIO or PTI. NO2 · also reacts with NO · to form N2O3, which is hydrolyzed to nitrite.
Alternative explanations for the cell cycle perturbations and apoptosis after NO · depletion may be related to the NO · targets guanylyl cyclase and cGMP. In support of this, exposure of RLE cells to the guanylyl cyclase inhibitor ODQ mimicked the effects observed with PTIO. Furthermore, 8-BrcGMP but not 8-BrcAMP ameliorated the cell cycle perturbations observed after exposure to PTIO or ODQ. Moreover, others (35) have reported that the addition of an NO · donor to endothelial cells could confer protection against tumor necrosis factor-induced apoptosis via a mechanism that involves a cGMP-dependent pathway. Our results demonstrate the involvement of endogenous NO · and cGMP-dependent pathways in the protection of lung epithelial cells from cell cycle perturbations and apoptosis. These data suggest that guanylyl cyclase and cGMP are downstream effectors of NO · that control cell cycle progression and apoptosis. To date, the targets of guanylyl cyclase and cGMP that control this phenotypic outcome in pulmonary cells are not known.
Numerous reports (14, 22, 30, 31, 39, 44) have already demonstrated that NO · and cGMP play an important role in the control of the cell cycle and apoptosis. For instance, NO · donors or cGMP-elevating agents have been shown to inhibit proliferation, block cell cycle progression, induce apoptosis (14, 22, 23, 30, 31, 39), and interfere with the epidermal growth factor-activated signal transduction pathway (44). Activities of cell cycle regulatory proteins including cyclin-dependent kinase-2 and phosphorylation of the retinoblastoma target protein were inhibited after exposure to an NO · donor (22). Induction of the cyclin-dependent kinase inhibitor p21 may play a role in the inhibition of cell cycle progression (22). An independent study (31) demonstrated that the arrest of cell cycle progression via inhibition of the cell cycle regulatory proteins, proliferating cell nuclear antigen, or cell division cycle 2 kinase is associated with increases of NO · release, further illustrating an inverse correlation between NO · and cell cycle progression. However, most of these studies use NO · donors to elevate NO · levels and do not assess the role of endogenous NO ·. Here we demonstrate that trapping of NO · from epithelial or mesothelial cells with PTIO also causes a block of cell cycle progression, characterized by decreases in BrdU incorporation and a block of cells in the late S and G2/M phases of the cell cycle. The findings suggest that NO · is required for completion of the cell cycle and that increases or decreases in the levels of NO · interfere with normal cell cycle progression.
The relationship between cell cycle perturbations and the apoptotic response observed after trapping of NO · is unclear. Conceivably, a block in cell cycle progression may be the trigger for apoptosis via the action of p53 and other cell cycle checkpoint regulators. Accumulation of p53 has been documented in conditions where NO · is being produced and is associated with apoptosis (8). Alternatively, apoptosis may result in compensatory proliferation of neighboring cells to repopulate injured epithelium, processes that occur in vivo in response to acute injury (4). However, this scenario is unlikely on the basis of the results in Fig. 4, where increases in the S and G2/M phases are observed before the induction of apoptosis.
In summary, our present findings demonstrate that depletion of
endogenous NO · causes cell cycle perturbations and death, possibly due to increased
H2O2
production or the altered function of cGMP-dependent processes. A
critical balance between O2· and
NO · may thus be important for cell cycle progression and survival of pulmonary cells.
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ACKNOWLEDGEMENTS |
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We appreciate the editorial comments of Prof. Brooke T. Mossman (Department of Pathology, University of Vermont, Burlington). We also thank Colette Charland for the flow cytometric analyses, Dr. Douglas Taatjes (Cell Imaging Facility, University of Vermont) for assistance with confocal microscopy, Dr. Victor Darley-Usmar for valuable advice, and Laurie Sabens for preparing the manuscript.
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FOOTNOTES |
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This work was supported by National Institute of Occupational Safety and Health Grant R03-OH-03467-01.
Y. M. W. Janssen is a Fellow of the Parker B. Francis Foundation for Pulmonary Research.
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. §1734 solely to indicate this fact.
Address for reprint requests: Y. M. W. Janssen, Dept. of Pathology, Univ. of Vermont, Medical Alumni Bldg., Burlington, VT 05405.
Received 22 May 1998; accepted in final form 18 September 1998.
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