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American Journal of Respiratory Cell and Molecular Biology. Vol. 28, pp. 443-450, 2003
© 2003 American Thoracic Society
DOI: 10.1165/rcmb.2002-0153OC

Hyperoxia Impairs Antibacterial Function of Macrophages Through Effects on Actin

Philip J. O'Reilly, Judy M. Hickman-Davis, Ian C. Davis and Sadis Matalon

Department of Medicine, Division of Pulmonary and Critical Care Medicine; Department of Anesthesiology; Department of Genomics and Pathobiology; and Department of Physiology and Biophysics, School of Medicine, University of Alabama at Birmingham, Birmingham, Alabama

Address correspondence to: Dr. Sadis Matalon, Dept. of Anesthesiology, University of Alabama at Birmingham, 901 19th Street South, BMR II, Rm. 224, Birmingham AL 35205-3703. E-mail: Sadis.Matalon{at}ccc.uab.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Oxidative stress may impair alveolar macrophage function in patients with inflammatory lung diseases or those exposed to high concentrations of oxygen. We investigated putative mechanisms of injury to macrophages by oxidative stress, using RAW 264.7 cells exposed to 95% oxygen for 48 h. Hyperoxia-exposed macrophages were less able to phagocytose and kill Klebsiella pneumoniae than normoxic controls, despite increased production of nitric oxide, a free radical important in pathogen killing. Exposure of macrophages to hyperoxia had marked effects on the actin cytoskeleton, including increased actin polymerization, loss of cortical actin, formation of stress fibers, de novo synthesis of actin, and actin oxidation. Hyperoxia induced changes in cell morphology, with increased cell size and pseudopod formation. Exposure of macrophages to jasplakinolide, an agent that increases actin polymerization, also impaired their ability to phagocytose Klebsiella. Alveolar macrophages isolated from mice exposed to 100% oxygen for 84 h also demonstrated impaired phagocytic function, as well as similar effects on the actin cytoskeleton and cell morphology to macrophages exposed to hyperoxia in vitro. We conclude that oxidative stress in vitro and in vivo impairs macrophage antibacterial function through effects on actin.

Abbreviations: alveolar macrophages, AM • acute respiratory distress syndrome, ARDS • adenosine triphosphate, ATP • Dulbecco's modified Eagle's medium, DMEM • 2,4-dinitrophenylhydrazine, DNPH • interferon-{gamma}, IFN-{gamma} • jasplakinolide, JP • lipopolysaccharide, LPS • mitogen-activated protein kinase, MAPK • nitric oxide, NO • polyacrylamide gel electrophoresis, PAGE • phosphate-buffered saline, PBS • reactive oxygen species, ROS • sodium dodecyl sulfate, SDS • trifluoroacetic acid, TFA • tris (hydroxymethyl) aminomethane, TRIS


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Hyperoxia and other forms of oxidative stress are known to be toxic to cells. The toxic effects of hyperoxia may be caused by increased intracellular generation of reactive oxygen species (ROS) (1) and may be mitigated by administration of antioxidants and overexpression of protective genes (2, 3). ROS may cause injury to cells through effects on macromolecules, including lipid peroxidation, modification of amino acid residues and nucleotides, and induction of nucleic acid strand breaks (4), or through activation of signal transduction cascades, such as the mitogen-activated protein kinase (MAPK) pathway (5, 6).

Hyperoxia can induce apoptosis or necrosis in cells, and arrests cell proliferation in different phases of the cell cycle, depending on the cell type (7–11). It can impair adenosine triphosphate (ATP) production through inhibition of iron-containing respiratory enzymes such as aconitase (12). Hyperoxia and ROS have been described to cause increased permeability of endothelial and epithelial monolayers (13, 14), and the exposure of animals to hyperoxia causes acute lung injury (injury to the alveolar epithelium and capillary endothelium, permeability pulmonary edema, and death [8, 15]).

In humans, the lung is exposed to ROS from environmental pollutants and during administration of high concentrations of oxygen to critically ill patients and patients with respiratory diseases. During pulmonary inflammation, infiltrating neutrophils and other inflammatory cells generate high concentrations of ROS. In the acute respiratory distress syndrome (ARDS), damage induced by neutrophil-generated ROS may be exacerbated by the administration of high concentrations of oxygen.

We hypothesize that hyperoxia and ROS impair lung innate immune function in patients with respiratory diseases such as ARDS, increasing their risk for pneumonia and death. Indeed, ARDS patients are more likely to develop ventilator-associated pneumonia than are other ventilated patients (16, 17), and are most likely to die from sepsis and the accompanying multi-organ dysfunction syndrome (18, 19). Alveolar macrophages (AM) play a critical role in lung innate immunity by phagocytosing and killing pathogens through the production of ROS and nitric oxide (NO) (20). Macrophages exposed to hyperoxia in vitro and AM isolated from hyperoxia-exposed animals exhibit impairment of chemotaxis, adherence, phagocytosis, pathogen-killing, and other functions (21–24). However, little information is available on how these effects on macrophage function are mediated.

In these experiments, we examined putative mechanisms of injury to macrophages from oxidative stress. Isolation of sufficient AM from ARDS patients for mechanistic studies is difficult due to the acuity of illness and contamination with inflammatory cells. Accordingly, we used RAW 264.7 cells, a murine macrophage cell line, exposed to hyperoxia in vitro, and AM isolated from hyperoxia-exposed mice. We suggest that our findings may be of relevance to patients with ARDS or to those receiving high concentrations of supplemental oxygen.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
All chemicals were from Sigma (St. Louis, MO) unless otherwise specified.

Cell Culture
RAW 264.7 cells (Cat. #CRL-2278; ATCC, Manassas, VA) were used in experiments. RAW cells were cultured in Dulbecco's modified Eagle's medium (DMEM) without phenol red (Cellgro, Herndon, VA) with 10% fetal bovine serum and 1% penicillin/streptomycin in 10-cm culture dishes and 75-cm2 culture flasks in 5% CO2 (normoxia) for 24 h until subconfluent. Selected cultures were then moved to sealed humidified chambers containing 95% O2/5% CO2 (hyperoxia) and incubated at 37°C for a further 48 h. RAW cells were resuspended by scraping and used in experiments. Cells were counted using a hemacytometer and viability determined using trypan blue exclusion. Apoptotic cells were detected by binding of annexin V-FITC (Pharmingen, San Diego, CA) and enumerated by flow cytometry (FACScan; Becton-Dickinson Inc., Franklin Lakes, NJ) using CELLQuest software. For cell cycle analysis, cells were fixed with 75% ethanol in 50 mM phosphate-buffered saline (PBS), stained with 10 µg propidium iodide in 1.1% sodium citrate with 1 mg/ml RNase A and analyzed by flow cytometry.

Bacterial-Killing Assay
RAW cells were activated with 100 U/ml recombinant murine interferon (IFN)-{gamma} (Pierce, Rockford, IL) and 0.5 µg/ml Escherichia coli lipopolysaccharide (LPS) for 4 h, and resuspended in serum-free DMEM in 5 ml polypropylene vials at 2 x 106 cells per vial. Klebsiella pneumoniae (ATCC 43816, type 2) were grown to log phase in brain heart infusion broth (Becton-Dickinson Inc., Sparks, MD), washed and resuspended in DMEM, and added to the vials at a multiplicity of infection of ~ 10:1 bacteria per cell. Final reaction volume was 0.5 ml. Infected cells were incubated at 37°C on a shaker for 30 min, washed three times by centrifugation to remove unattached bacteria, and resuspended in 0.5 ml DMEM (25). A 100-µl aliquot was taken at this point (0 min), macrophages lysed by sonication, and viable bacteria enumerated by serial dilution and culture on nutrient agar plates (Becton-Dickinson Inc.). Macrophages and bacteria were reincubated for a further 60 min, at which point another 100-µl aliquot was taken and viable bacteria determined as before.

Measurement of NO Production by Macrophages
RAW cells were resuspended in serum-free DMEM, seeded in 24-well culture plates at 7.5 x 105 cells per well and incubated with 100 U/ml IFN-{gamma} and 0.5 µg/ml LPS for 6 h. NO production was quantified by measuring the levels of nitrite and nitrate, the stable oxidation products of NO, in the cell supernatants using the Greiss reaction. Nitrate was first converted to nitrite by incubation with E. coli reductase for 1 h at 37°C. One hundred microliters of sample was then incubated in duplicate with equal volumes of 1% sulfanilamide and 0.1% N-1-naphthylethylene-diamine dihydrochloride for 10 min, and the absorbance read at 550 nm on a spectrophotometer (26).

Phagocytosis Assay
K. pneumoniae were labeled with fluorescein, as described (26) and stored in 10% glycerol in dH2O at -20°C. RAW cells (5 x 105) were incubated with fluorescent bacteria in 1 ml serum-free DMEM on a shaker at 37°C and 5% CO2 for 1 h, pelleted by centrifugation at 500 x g, and washed three times in ice-cold PBS with 0.02% EDTA to stop phagocytosis and remove extracellular bacteria. Cells were incubated for 5 min in 0.02% trypan blue in PBS to quench extracellular fluorescence (27), resuspended in ice cold PBS/EDTA and analyzed by flow cytometry. RAW cells were identified based on forward and side scatter characteristics and ten thousand events per sample were analyzed. Phagocytosis was measured using the increase in mean cell fluorescence of macrophages after incubation with bacteria (26).

ATP Measurement
ATP was measured with a kit (FL-ASC, Sigma, St. Louis, MO). Macrophages were seeded 4 x 104 cells per well in a 96 well microplate (Costar, Corning, NY). Released ATP reacted with luciferin and firefly luciferase to produce light which was detected with a luminometer (Lumistar; B&L Systems, Maarssen, Netherlands). ATP was quantified using an ATP standard.

Actin Western Blot
RAW cells were lysed in RIPA lysis buffer (150 mM NaCl, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate [SDS], 50 mM tris [hydroxymethyl] aminomethane [TRIS], pH 8.0), containing a protease inhibitor cocktail (Complete Mini; Roche, Mannheim, Germany). Protein concentrations of the whole cell lysates were measured using the bicinchoninic acid protein assay (Pierce) and equal amounts of protein per sample were resolved by 8% SDS-polyacrylamide gel electrophoresis (PAGE) and transferred to a polyvinylidene fluoride (PVDF) membrane. Membranes were probed with a monoclonal anti-actin antibody (0.4 µg/ml; RDI, Flanders, NJ), followed by a horseradish peroxidase (HRP)-conjugated secondary (1:10,000). Immunoreactive protein complexes were detected by enhanced chemiluminescence.

Actin Immunoprecipitation and Western Blot Detection of Carbonyl Formation
RAW cells were lysed by sonication in a hypotonic lysis buffer (50 mM NaCl, 50 µM MgCl2, 5 mM EGTA, 0.1% Triton X-100, 50 mM TRIS, pH 7.4), containing 50 mM DTT (to prevent protein oxidation) and protease inhibitors, and incubated with 20 µM cytochalasin D for 1 h on ice with mixing. Cell lysates were cleared by centrifugation at 16,000 x g and the supernatants incubated overnight with 20 µl/ml Protein A/G Plus Agarose (Santa Cruz Biotechnology, Santa Cruz, CA) and 2 µg/ml polyclonal goat anti-actin (RDI) at 4°C with rotation. Immunoprecipitates were washed three times in wash buffer (300 mM NaCl, 0.1% Triton X-100, 50 mM TRIS, pH 7.4), once in wash buffer plus 0.1% SDS and 0.1% sodium deoxycholic acid, and once in PBS. Immunoprecipitates were resuspended in 6% SDS and protein concentrations determined using the bicinchonic acid assay. Samples were incubated with one volume 20 mM 2,4-dinitrophenylhydrazine (DNPH) in 10% trifluoroacetic acid (TFA) for 20 min at room temperature to convert carbonyls to hydrazone derivatives, and the reaction was terminated by the addition of 0.75 vol 2M TRIS/30% glycerol/19% ß-mercaptoethanol (28). Controls were incubated with TFA alone without DNPH. Equal amounts of protein per sample were resolved by 8% SDS-PAGE and transferred to a PVDF membrane. Carbonyls were detected with a primary rabbit anti-dinitrophenyl (DNP) antibody and HRP-conjugated secondary (Oxyblot, Intergen, Purchase, NY) according to the manufacturer's instructions.

Immunofluorescence Microscopy and F-Actin Quantitation
RAW cells were adhered to glass coverslips for 2 h, fixed with 3% formaldehyde, permeabilized with 0.1% Triton X-100 in PBS, and stained with Texas Red X-phalloidin (Molecular Probes, Eugene, OR), diluted 1:40 in 1% bovine serum albumin. Nuclei were counterstained with 20 µg/ml Hoechst 33258 in PBS. Coverslips were mounted in 0.1% para-phenylenediamine in 90% glycerol and sealed with nail polish. Images were obtained with a Leitz Orthoplan microscope (Leica, Wetzlar, Germany) using IPLab software (Scanalytics, Fairfax, VA). For actin quantitation, 4 x 105 RAW cells were seeded in a 96-well microplate (Greiner, Frickenhausen, Germany) for 2 h, fixed, permeabilized, and stained with 0.5 µg/ml Hoechst 33258 and Texas Red X-phalloidin diluted 1:2 in PBS for 45 min. Monolayers were washed three times in PBS and analyzed on a SpectraMax Gemini fluorometer (Molecular Devices, Sunnyvale, CA) with dual excitation and emission wavelengths for Hoechst 33258 and Texas Red X. Average cell F-actin content was determined by comparing phalloidin and DNA staining in individual wells, as described previously (29).

Exposure of Mice to Hyperoxia and Isolation of AM
Male C57BL/6NCr mice were obtained from Frederick Cancer Research and Development Center (National Cancer Institute, Frederick, MD), maintained in autoclaved microisolator cages (Lab Products, Maywood, NJ), and provided with food (Teklad, Madison, WI) and water ad libitum. All mice used in studies were 6–8 wk old and 20–25 g in weight. Mice were exposed to 100% O2 for 84 h (sublethal hyperoxia) in a specially designed chamber, as previously described (30). Concentrations of O2 were maintained above 99%, measured with an O2 analyzer (Ohmeda 5120; Ohmeda Medical, Madison, WI). After exposure, mice were killed with an intraperitoneal injection of ketamine–xylazine, and a sterile 18-gauge intravenous catheter was inserted caudally into the lumen of the exposed trachea. The lungs were lavaged in situ with 10 separate 1-ml aliquots of sterile saline. Bronchoalveolar lavage fluid was centrifuged and the cellular pellet resuspended in DMEM. AM were used for phagocytosis, using fluorescent Klebsiella as before, or adhered to Lab-Tek chamber slides (Nunc, Naperville, IL) and stained with Texas Red X-phalloidin for F actin quantitation. Slides were mounted with Vectashield with DAPI (Vector Laboratories, Burlingame, CA) and examined by immunofluorescence microscopy using the x25 objective. Fifteen macrophages were randomly selected per slide and analyzed for total and mean phalloidin fluorescence and for cell size, using IPLab software.

Statistical Analysis
Experiments were performed three times in triplicate except where specified. Results are expressed as mean ± SEM. Data were analyzed using a two-sample t test for parametric data or Wilcoxon Rank Sum test for nonparametric data with Statistix 7 (Analytical Software, Tallahassee, FL). A P value <= 0.05 was considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Hyperoxia Alters Cell Morphology and Induces Cell Cycle Arrest and Apoptosis in RAW 264.7 Macrophages
We found that culture in 95% O2 greatly reduced numbers of RAW cells (Figure 1). Propidium iodide staining revealed that hyperoxia induced growth arrest in G0/G1 phase of the cell cycle (71 ± 2% of hyperoxic cells in G0/G1 versus 52 ± 2% of normoxic cells, n = 6, P < 0.05). Hyperoxia also caused a small increase in the percentage of RAW cells in early apoptosis as detected by annexin binding (5.9 ± 1% of hyperoxic cells versus 1.4 ± 0.35% of normoxic cells, n = 3, P < 0.05%). RAW cells cultured in hyperoxia also appeared larger when examined under light microscopy and had altered morphology with increased pseudopod formation (Figure 1).



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  		Figure 1. Hyperoxia reduces proliferation and alters cell morphology in RAW 264.7 cells. A quantity of 3 x 106 RAW cells was cultured in 21% O2 for 24 h (normoxia), after which selected cultures were moved to 95% O2 (hyperoxia). After a further 48 h, images were taken of cell monolayers cultured in normoxia (A) or hyperoxia (B) using an Olympus IX 70 inverted microscope and UltraView software (Perkin-Elmer, Wellesley, MA).

 
Hyperoxia Reduces the Antibacterial Properties of RAW 264.7 Macrophages
RAW cells cultured in hyperoxia and activated with LPS and IFN-{gamma} demonstrated an impaired ability to kill K. pneumoniae compared with controls cultured in normoxia, as illustrated by increased proliferation of bacteria (Figure 2A). However, when stimulated with LPS and IFN-{gamma}, hyperoxic RAW cells produced significantly greater amounts of NO than normoxic controls (Figure 2B). This makes it unlikely that the impaired killing of Klebsiella was due to an inability to produce NO. We then investigated the ability of hyperoxic RAW cells to phagocytose pathogens by incubating them with fluorescent Klebsiella. Cells were incubated with trypan blue to quench extracellular fluorescence and analyzed by flow cytometry (Figure 2C). Phagocytosis was quantified using the increase in mean cell fluorescence over baseline (autofluorescence) after incubation with bacteria. Phagocytosis by hyperoxic macrophages was expressed as a percentage of that by normoxic macrophages examined simultaneously. We found that hyperoxic macrophages phagocytosed significantly fewer Klebsiella than normoxic controls (63 ± 9%, n = 9, P < 0.05). This provides a possible mechanism for the impaired pathogen killing observed in RAW cells exposed to hyperoxia. In accordance with their morphology under light microscopy (Figure 1), hyperoxic RAW cells were larger than normoxic controls when analyzed by flow cytometry (greater forward scatter), and also had greater autofluorescence (data not shown).



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  		Figure 2. (A) Hyperoxia reduces pathogen killing by RAW 264.7 cells. RAW cells were cultured in 21% (black bars) or 95% O2 (hatched bars), activated with IFN-{gamma} and LPS, and incubated with K. pneumoniae to allow adherence/phagocytosis. Macrophages were washed to remove free bacteria and incubated for 60 min. Aliquots were taken at 0 and 60 min, and bacteria quantified by serial dilution and culture. Data represent three separate experiments performed in triplicate. *P < 0.05 comparing normoxic and hyperoxic macrophages at 60 min. cfu, colony-forming units. (B) Hyperoxia increases NO production by RAW 264.7 cells. RAW cells cultured in 21% (black bars) or 95% O2 (hatched bars) were stimulated with IFN-{gamma} and LPS or not (null). Total nitrite and nitrate was measured in the supernatant after 6 h and expressed as nmol NO per 106 cells per hour. Data represent three separate experiments performed in triplicate. *P < 0.05 comparing normoxic and hyperoxic macrophages. (C) Hyperoxia reduces phagocytosis by RAW 264.7 cells. RAW cells cultured in 21% or 95% O2 were incubated with fluorescent K. pneumoniae, washed, and incubated with 0.02% trypan blue to quench extracellular fluorescence, and then analyzed by flow cytometry. Two populations of macrophages were identified, N (nonphagocytosing) and P (phagocytosing), according to fluorescence intensity, measured in fluorescence units and plotted on the x-axis.

 
Hyperoxia Impairs Phagocytosis by RAW 264.7 Macrophages through Effects on Actin Polymerization
We investigated possible mechanisms of impairment of phagocytosis in RAW cells exposed to hyperoxia. Phagocytosis depends on the directional polymerization of actin to surround and internalize the pathogen and is an energy-dependent process. We found a small but nonsignificant increase in ATP content of hyperoxic RAW cells compared with normoxic controls (10.2 ± 1.45 versus 8.1 ± 0.77 pg per cell, n = 5, P = 0.23). Thus, we believe that impaired phagocytosis by macrophages cultured in hyperoxia was not due to an effect on metabolism limiting the availability of ATP. Using quantitative Western blotting, we found that hyperoxic RAW cells contained increased amounts of actin compared with normoxic controls (Figure 3), implying that impaired phagocytosis by macrophages cultured in hyperoxia was not due to reduced availability of actin. However, when we examined the actin cytoskeleton by fluorescence microscopy using phalloidin, a specific probe for polymerized (F) actin, we found marked changes in hyperoxia-exposed RAW cells compared with normoxic controls. Macrophages cultured in normoxia demonstrated the cortical distribution of F actin normally seen in quiescent cells (Figure 4A). In contrast, macrophages cultured in hyperoxia demonstrated loss of cortical actin, increased formation of stress filaments and aggregates of actin, and an apparent overall increase in actin polymerization (Figure 4B). When F actin content in RAW cells was quantified using a well characterized fluorometric assay, we confirmed that actin polymerization was increased approximately 2-fold in hyperoxic macrophages compared with normoxic controls (Figure 4C).



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  		Figure 3. Hyperoxia increases total actin in RAW 264.7 cells. RAW cells cultured in 21% or 95% O2 were lysed and equal amounts of protein resolved using SDS-PAGE and transferred to a PVDF membrane. The membrane was probed using a monoclonal anti-actin antibody, and developed using enhanced chemiluminescence to reveal a 43-kD band corresponding to monomeric actin. Figure is representative of two separate experiments.

 


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  		Figure 4. Hyperoxia modifies polymerized actin in RAW 264.7 cells. RAW cells cultured in 21% or 95% O2 were fixed, permeabilized, and stained with phalloidin conjugated to Texas Red X, and with Hoechst 33258. Normoxic cells (A) demonstrated a normal cortical distribution of polymerized actin, whereas hyperoxic cells (B) demonstrated increased amounts of polymerized actin, with formation of stress filaments and loss of cortical actin. For actin quantitation (C), RAW cells were stained with Hoechst 33258 and Texas Red X-phalloidin, and analyzed using a microplate fluorometer with dual excitation and emission wavelengths for both fluorochromes. The ratio of Texas Red X staining to that of Hoechst 33258 was used to estimate the amount of polymerized actin per cell. Data represent three experiments performed in triplicate. *P < 0.05 comparing normoxic cells (black bar) to hyperoxic cells (hatched bar)

 
To determine whether increased polymerization of actin might account for the impairment of phagocytosis by hyperoxia, we exposed macrophages to jasplakinolide (JP), a macrocyclic cyclodepsipeptide and analog of phalloidin. Unlike phalloidin, JP can permeate living cells, where it increases and stabilizes polymerized actin (31). RAW 264.7 cells were exposed to 1 µM JP for 1 h and examined for their ability to phagocytose fluorescent Klebsiella. Phagocytosis was measured using the increase in mean cell fluorescence over baseline, and expressed as units of fluorescence. Phagocytosis by JP-exposed RAW cells was significantly reduced compared with phagocytosis by RAW cells exposed to vehicle alone (dimethyl sulfoxide) (9.16 ± 0.72 versus 4.34 ± 0.37, n = 9, P < 0.05). Exposure to JP had no effect on cell viability (data not shown). This indicates that increased actin polymerization is a potential mechanism of impaired phagocytosis in macrophages exposed to hyperoxia.

Hyperoxia Induces Oxidation of Actin in RAW 264.7 Cells
We considered that the effects of hyperoxia on actin polymerization and phagocytosis might result from chemical modification of amino acid residues in actin by ROS generated intracellularly. To investigate this, we used SDS-PAGE and Western blotting of immunoprecipitated actin from RAW cells cultured in normoxia or hyperoxia to detect carbonyl formation as a marker of amino acid oxidation. Actin from hyperoxic macrophages demonstrated significant immunoreactivity for carbonyl groups, whereas actin from normoxic macrophages demonstrated no increase in carbonyl reactivity over its own negative control (Figure 5). This indicates that hyperoxia induces oxidation of actin in RAW 264.7 cells. However, when Western blots were probed for nitrotyrosine, for evidence of actin nitration, no immunoreactivity of immunoprecipitated actin was detectable following culture in hyperoxia (data not shown).



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  		Figure 5. Hyperoxia induces oxidation of actin in RAW 264.7 cells. Actin was immunoprecipitated from RAW cells cultured in 21% or 95% O2 and incubated with 2,4-DNPH in TFA to convert carbonyl groups to their 2,4-DNP–hydrazone derivatives, or with TFA alone as a negative control. Equal amounts of protein per lanewere resolved on SDS-PAGE and Western blotting performed with an anti-DNP primary and an HRP-conjugated secondary antibody. Derivatized samples (A and C) were compared with their own negative controls (B and D) to control for nonspecific binding. Actin from hyperoxic macrophages (43 kD) demonstrated strong immunostaining for carbonyl group formation (C) compared with its negative control (D), whereas actin from normoxic macrophages (A) demonstrated no increased staining over control (B). Figure is representative of three separate experiments.

 
AM Isolated from Mice Exposed to 100% O2 In Vivo Demonstrate Impaired Phagocytic Function, Increased Actin Polymerization, and Changes in Morphology
We exposed C57BL/6NCr mice to sublethal hyperoxia (100% O2 for 84 h), which did not cause any mortality, and performed bronchoalveolar lavage to obtain AM. Cells obtained by bronchoalveolar lavage were at least 90% AM when examined by Diff-Quik (Dade-Behring, Newark, DE) staining of cytospin preparations (data not shown). We examined the ability of AM to phagocytose fluorescent Klebsiella as before. AM from six hyperoxia-exposed mice demonstrated an almost 30% decrease in phagocytic function compared with AM from six normoxic controls (Figures 6A and 6B).



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  		Figure 6. AM from hyperoxia-exposed mice demonstrate impaired phagocytosis. AM from hyperoxia-exposed mice were examined for their ability to phagocytose fluorescent Klebsiella, using flow cytometry as with RAW cells, except that 3 x 104 AM were used per assay. Phagocytosis was measured using the increase in mean cell fluorescence of AM after incubation with bacteria and expressed as fluorescence units (x-axis) (A). AM from hyperoxic mice (hatched bar) demonstrated significantly impaired phagocytosis when compared with AM from normal controls (black bar) (B). Figure represents data from six mice per group with experiments performed in triplicate. *P < 0.05.

 
AM from eight hyperoxia-exposed mice and seven controls were examined by fluorescence microscopy for cell size and F actin content. Data from hyperoxic mice were averaged and expressed as a percentage of those from control mice, whose AM were harvested and examined simultaneously. We found that AM from hyperoxic mice were significantly larger (126 ± 2%, n = 8, P < 0.05) than AM from controls, and demonstrated increased total staining for polymerized actin (134 ± 10%, n = 8, P < 0.05). There was no significant difference in mean staining for polymerized actin between AM from hyperoxic mice and normoxic controls (106 ± 8%, n = 8). These data are similar to those obtained from RAW 264.7 cells exposed to hyperoxia in vitro, which also demonstrated impaired phagocytosis, increased actin polymerization, and increased cell size compared with controls.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Potential injury to lung innate immunity from ROS is of great importance in the management of patients with respiratory disease, increasing their susceptibility to pneumonia and death. Surfactant protein-A, a protein important in lung immunity, is nitrated in ARDS (32), which impairs its ability to mediate phagocytosis and killing of pathogens (33). It is likely that similar injury occurs to AM, another important component of lung immunity. Functional defects have been demonstrated in AM isolated from animal models of ARDS (34, 35) and macrophages exposed to hyperoxia (21–24). However, there is, as yet, little mechanistic information on how injury to macrophages from ROS occurs.

To address this question, we initially performed a series of experiments using RAW 264.7 macrophages, exposed to 95% O2. We found that hyperoxia induced apoptosis and cell cycle arrest in G0/G1 phase in RAW 264.7 macrophages. Hyperoxia is well known to be toxic to cultured cells, but the mechanism of cell death (apoptosis versus necrosis) and phase of the cell cycle in which growth arrest occurs differ depending on the cell type involved (7, 9, 10). Hyperoxia-induced apoptosis in RAW 264.7 cells has been reported to be mediated via the ERK p42/p44 MAPK pathway (6).

We demonstrate that RAW 264.7 cells cultured in hyperoxia are less able than control cells, cultured in normoxia, to phagocytose and kill K. pneumoniae, a gram-negative bacterium and common cause of nosocomial infections. These findings are consistent with previous reports of macrophages exposed to hyperoxia (21–24). Hyperoxia has also been shown to inhibit superoxide production and stimulate inflammatory cytokine production by macrophages (24, 36–38). It appears that macrophage cell lines, such as RAW 264.7, are less effective at killing pathogens than primary macrophages (39). Consistent with this, we found that Klebsiella numbers increased during the course of the killing assay, but proliferation of bacteria was greater when they were incubated with hyperoxic macrophages.

To further investigate the mechanism for impaired killing by hyperoxic macrophages, we measured their ability to produce NO, a free radical important in pathogen killing, and found that they produced considerably more NO, after stimulation with LPS and IFN-{gamma}, than did normoxic controls. These data are consistent with a recent study demonstrating increased NO production by rat AM exposed to hyperoxia for 24 h, mediated by intracellular generation of ROS and the transcription factors nuclear factor-{kappa}B and AP-1 (40). Thus, it seems unlikely that impaired killing by hyperoxic RAW 264.7 cells was due to reduced production of reactive nitrogen species.

The process of phagocytosis requires the polymerization of actin in a directional fashion to surround and internalize the target (41). Actin polymerization is dependent on adequate supplies of ATP. Exposure to hyperoxia can impair the activity of cellular enzymes important in oxidative metabolism (12, 42), which could lead to increased anaerobic glycolysis, depletion of glucose in the cell growth medium, and subsequent depletion of ATP (43). However, ATP levels in RAW 264.7 cells cultured in hyperoxia were not reduced. Glucose content in the cell growth medium was higher in the hyperoxic cultures, likely as a result of inhibition of cell proliferation by hyperoxia (data not shown).

Examination of the actin cytoskeleton of RAW 264.7 cells cultured in hyperoxia, by immunofluorescence microscopy, revealed marked changes, including an increase in the degree of actin polymerization, loss of cortical actin, and the formation of prominent stress fibers and actin aggregates. Quantitative Western blotting revealed that the amount of actin in RAW 264.7 cells exposed to hyperoxia was increased compared with normoxic controls, implying that de novo synthesis of actin occurred, perhaps secondary to the depletion of actin monomers. Exposure of RAW 264.7 macrophages to JP, a cell permeant analog of phalloidin that increases and stabilizes polymerized actin in living cells (31), reduced the ability of RAW 264.7 macrophages to phagocytose fluorescent Klebsiella by ~ 50%. This indicates that increased actin polymerization is a potential mechanism explaining impairment of phagocytosis by hyperoxic RAW cells.

Exposure of endothelial and epithelial cells to ROS has been shown to modify (increase or decrease) actin polymerization, disrupt actin filaments, and induce loss of cortical actin, formation of stress fibers, and de novo synthesis of actin (5, 44–46). These effects are believed to be important in mediating permeability changes in inflammatory diseases (47, 48). As far as we are aware, our studies are the first to describe similar effects and their functional significance in macrophages. Indeed, modification of actin could affect numerous cellular functions, including ion transport across epithelia (49), receptor trafficking to cell membranes, and the activity of enzymes such as inducible nitric oxide synthase and NADPH oxidase (50, 51).

We found that exposure of mice to 100% O2 for 84 h impaired phagocytic function and increased actin polymerization and cell size in their AM. These effects are similar to those in RAW 264.7 cells exposed to hyperoxia in vitro. As far as we are aware, this is the first demonstration of the effects of ROS on actin polymerization in an in vivo model. Effects on actin may have been due to ROS produced by inflammatory cells, recruited to the lung as a result of hyperoxia-induced injury, or to a direct effect of the hyperoxia itself. However, very few neutrophils were present in the bronchoalveolar lavage fluid of these mice (data not shown), suggesting that the effects observed were predominantly due to hyperoxia.

The mechanism whereby hyperoxia exerts these effects on actin is unclear. Effects of hyperoxia on actin polymerization in endothelial cells have been linked to depletion of ATP (52), but this does not appear to be the mechanism in RAW 264.7 cells. NO has been reported to increase actin polymerization and pseudopod formation in RAW 264.7 cells (53) and to inhibit pseudopod formation and phagocytosis in mouse peritoneal macrophages (54). NO can also combine with superoxide to produce reactive species which can nitrate and oxidize proteins (55). Nitration and oxidation of actin has been demonstrated in numerous cell types exposed to ROS and inflammatory cytokines (47, 56, 57). In all of these cases, the effects on actin were dependent on NO generated by inducible nitric oxide synthase within the cells affected. This illustrates how NO, as well as fulfilling an important role in host defense, can be toxic to the cells that produce it (58).

We were able to detect actin oxidation but not nitration in RAW 264.7 cells exposed to hyperoxia, possibly as insufficient quantities of reactive oxygen and nitrogen species were generated by this stimulus. It is not clear how oxidation of actin leads to increased actin polymerization within cells. In fact, oxidation of actin monomers in a cell-free system reduces their ability to polymerize in vitro, which is in contrast to the effects seen here (59). Exposure of endothelial cells to hydrogen peroxide increased actin polymerization through activation of p38 MAPK and phosphorylation of heat shock protein 27 (5). The relevance of this to RAW 264.7 cells exposed to hyperoxia is uncertain, as p38 MAPK has not been reported to be activated in this case (6).

Although the concentrations of oxygen used in these experiments were high (95–100%), we believe that our findings are relevant to patients with respiratory diseases for a number of reasons. Studies of animals and macrophages exposed to concentrations of oxygen ranging from 40–85% have demonstrated similar effects on macrophage function at these lower levels, including impairment of phagocytosis and pathogen-killing and increased NO production (21–23, 40). Effects on the actin cytoskeleton have also been seen in cells exposed to lower levels of oxygen (45). Furthermore, it is not uncommon for patients with severe lung injury to be treated with concentrations of oxygen as high as 100% for prolonged periods. In addition, local concentrations of oxidant molecules, such as hydrogen peroxide, superoxide and peroxynitrite, may be very high in the alveolar space in inflammatory lung diseases. Therefore, we believe that the effects of 95–100% oxygen that we describe on macrophage function are relevant to the clinical setting.

In summary, oxidative stress induces changes in the polymerization state of actin in macrophages, which impairs their ability to phagocytose pathogens. This is of relevance to patients with ARDS and other inflammatory lung diseases and to patients treated with high concentrations of oxygen, and may, in part, explain the high incidence of pulmonary infections seen in these patients.


   Acknowledgments
 
This work was supported by grants HL31197, HL51173 P30, and DK54781 (S.M.) from the National Institutes of Health. J.M.H.-D. was supported by RR00149 and P.J.O. by a fellowship award from the American Heart Association (Southeastern affiliate). I.C.D. is a Parker B. Francis fellow. The authors would like to acknowledge the valuable input of Dr. K. Randall Young in this project and thank Ms. Glenda Davis, Ms. Carpantato Myles, and Dr. Kedar Shrestha for their excellent technical assistance with these studies.

Received in original form August 13, 2002

Received in final form October 14, 2002


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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