Logo of iaiPermissionsJournals.ASM.orgJournalIAI ArticleJournal InfoAuthorsReviewers
Infect Immun. 2008 Oct; 76(10): 4455–4462.
Published online 2008 Jul 21. doi:  10.1128/IAI.00136-08
PMCID: PMC2546817

Mycoplasma pneumoniae Infection and Environmental Tobacco Smoke Inhibit Lung Glutathione Adaptive Responses and Increase Oxidative Stress

Abstract

Chronic cigarette smoking evokes a lung glutathione (GSH) adaptive response that results in elevated GSH levels in the lung epithelial lining fluid (ELF). Currently, little is known about how the lung regulates or maintains steady-state levels of ELF GSH. Pathogens such as Mycoplasma pneumoniae can exacerbate airway inflammation and oxidative stress. The present study examined whether M. pneumoniae infections synergize with environmental tobacco smoke (ETS) to disrupt lung GSH adaptive responses. Mice were exposed separately and in combination to ETS and M. pneumoniae for 16 weeks. ETS exposure resulted in a doubling of ELF GSH levels, which was blocked in the M. pneumoniae-exposed mice. In addition, the ETS-plus-M. pneumoniae-exposed mice had elevated levels of oxidized glutathione (GSSG), resulting in a dramatic change in the ELF redox state that corresponded with an increase in lung tissue DNA oxidation. Similar findings were observed in human lung epithelial cells in vitro. Cells exposed separately or in combination to cigarette smoke extract and M. pneumoniae for 48 h had elevated apical levels of GSH compared to control cells, and these increases were blocked by M. pneumoniae and were also associated with increased cellular DNA oxidation. Further studies showed that M. pneumoniae exposure blocked ETS-induced increases in GSH reductase, an enzyme that recycles GSSG back to GSH, both in vitro and in vivo. These studies suggest that M. pneumoniae infection synergizes with ETS and suppresses the lung's ability to respond appropriately to environmental challenges leading to enhanced oxidative stress.

The lung's large surface area makes it susceptible to potential adverse effects of airborne environmental agents (13). Agents such as cigarette smoke, ozone, nitrogen oxides, and fuel emissions react with the lung epithelium to generate reactive species (RS) (17, 45). These RS can directly or indirectly damage lung proteins, lipids, and DNA, contributing to lung oxidative stress. Environmental tobacco smoke (ETS) contains more than 4,700 chemical entities, generating approximately 1014 RS per puff (36). These RS are found in both the gas and tar phases. Cigarette smoking is implicated as a primary risk factor for the development of many lung diseases, including chronic obstructive pulmonary disease (COPD) (38, 42) and cancer (48, 50). However, even in the presence of the chronic oxidative burden of chronic cigarette smoking, only 20% of smokers develop COPD. We speculate that lung antioxidant adaptive responses may be an important protective factor in the 80% of chronic smokers who do not develop COPD. Although it has been hypothesized that the imbalance between oxidants and antioxidants plays a prominent role in the development of COPD, only a limited number of studies have explored whether agents that exacerbate lung disease may affect the lung's ability to mount and sustain antioxidant adaptive responses.

The lung is continually exposed to mixtures of pollutants and to airborne pathogens such as Pseudomonas aeruginosa and Mycoplasma pneumoniae (26). Previous studies suggest that M. pneumoniae infection not only causes respiratory diseases but is also responsible for extrapulmonary complications (8). M. pneumoniae has been shown to increase oxidative stress in epithelial cells (1, 2, 21). In addition, M. pneumoniae is also known to increase COPD exacerbations, and a subset of COPD patients have chronic M. pneumoniae infections (32). M. pneumoniae is difficult to culture, which makes this organism hard to detect (14), and M. pneumoniae infections are likely underreported.

The lung is well defended against the deleterious effects of environmental oxidants by a number of potent antioxidants. The apical fluid bathing airway epithelium is also known as the epithelial lining fluid (ELF) and is the first to encounter these environmental agents. The ELF contains numerous enzymatic and nonenzymatic antioxidants, such as ascorbic acid, uric acid, glutathione (GSH), superoxide dismutase, catalase, GSH peroxidase, and plasma proteins (45). A major antioxidant found in the ELF is GSH. ELF GSH protects the lung by scavenging exogenously generated RS and is an essential cofactor for many cellular detoxification pathways. The normal concentration of GSH in the ELF is 10 to 100 times that found in the plasma (6). The cystic fibrosis transmembrane conductance regulator (CFTR) protein is currently the only known apical epithelial GSH transporter in the lungs but is responsible for regulating only 50% of basal ELF GSH levels (47).

The purpose of the current study was to examine the effect of potential interactions between ETS and M. pneumoniae infection on lung GSH adaptive responses. We report that ETS stimulates increases in the ELF GSH steady-state levels that are only partially dependent upon CFTR and that chronic ETS-mediated increases in GSH levels are impaired in M. pneumoniae-infected mice. Furthermore, our study also suggests that M. pneumoniae infections interfere with the lung's ability to keep the GSH in its reduced form and that this is associated with increased lung oxidative stress. We observed similar effects in cultured human lung epithelial cells, where M. pneumoniae infections blunted cigarette smoke extract (CSE)-induced increases in apical GSH levels. We have also discovered that M. pneumoniae infection inhibits ETS-induced GSH reductase (GR) levels, which correspond with elevated levels of oxidized GSH (GSSG) and increased markers of oxidative stress. These studies are the first to provide a mechanistic explanation for how M. pneumoniae may exacerbate lung disease in smokers by interfering with lung GSH adaptive responses and enhancing oxidative stress.

MATERIALS AND METHODS

Animals.

Pathogen-free 8- to 10-week-old BALB/c female mice were obtained from Jackson Laboratory (Bar Harbor, ME). Mice were fed Purina mouse chow 5010 and autoclaved tap water ad libitum. Mice were allowed to acclimate in the vivarium for at least 1 week before use. All animal procedures were approved by the IACUC committee at National Jewish Medical and Research Center.

M. pneumoniae preparation and inoculation.

Mycoplasma pneumoniae (strain FH; ATCC 15531) was cultured in SP-4 broth for approximately 10 days at 37°C, spun and resuspended in saline, and frozen in aliquots at −80°C. These aliquots were used for M. pneumoniae infections in cultured cells and BALB/c mice during the course of the experiment. Briefly, thawed M. pneumoniae aliquots were spun and resuspended in SP-4 broth and then incubated for 2 h at 37°C. The broth was collected and spun again at 6,000 rpm for 5 min. The resultant M. pneumoniae pellet was resuspended in sterile saline to yield approximately 1 × 108 CFU/50 μl.

BALB/c mice were anesthetized by administering 2,2,2-tribromoethanol (Aldrich, Milwaukee, WI) intraperitoneally (0.25 g/kg of body weight). Groups of 8 to 10 mice were inoculated intranasally with 50 μl of either saline or M. pneumoniae at approximately 1 × 108 CFU. After the inoculation, mice were placed at a 45° angle on their bedding for 5 min, allowing the M. pneumoniae to distribute evenly throughout the lungs. Mice were inoculated with M. pneumoniae 2, 10, and 16 weeks after the start of cigarette smoke exposure. Bronchoalveolar lavage (BAL) was performed 6 days after the last M. pneumoniae inoculation. The times and M. pneumoniae titers used in these studies were modeled from those in a previous study (11).

In vivo exposure to ETS.

Mice were exposed to ETS in a TE-10 smoking machine system (Teague Enterprises, Davis, CA). The TE-10 system is a microprocessor-controlled cigarette smoking system that produces both sidestream and mainstream smoke from filtered research cigarettes, which we refer to as ETS. Cigarettes (research grade 2RF4; University of Kentucky) are puffed using the Federal Trade Commission method of one 35-ml puff for 2 seconds once a minute. The sidestream smoke is collected in a chimney area above the burning cigarettes. The mainstream smoke is combined with the sidestream portion and used to maintain the smoke levels. In the mixing chamber, a fan homogenizes the vapor and particulate suspension, generating the ETS used in the exposure. All exposures were characterize for nicotine, carbon monoxide, and particulates. Nicotine levels were measured by drawing samples from the exposure chambers through sorbent tubes. Samples were extracted from the tubes, and nicotine levels were determined using gas chromatography. Carbon monoxide levels were monitored in the exposure chambers by a nondispersive infrared analyzer. Total suspended particulate levels in the exposure chambers were measured by using filter samples (Pallflex Products Co., Putnam, CT) and gravimetric analysis. The chambers were regulated to maintain an average total particulate concentration of 75 mg/m3, a carbon monoxide concentration of 190 ppm, and a nicotine concentration of 6 mg/m3. Mice were exposed to cigarette smoke for 6 h a day 5 days a week for 16 weeks. Plasma cotinine levels were measured by direct enzyme-linked immunosorbent assay (Immunalysis Corporation, Pomona, CA) in filtered air (FA)-, M. pneumoniae-, ETS-, and M. pneumoniae-plus-ETS-exposed mice and were 7.1, 8.3, 98.1 (58 to 100), and 88.9 (64.7 to 100), respectively (ng/ml; expressed as median values and ranges).

CSE preparation.

One Kentucky reference cigarette (research grade 2RF4; University of Kentucky) containing 26.8 mg tar and 2.45 mg nicotine was combusted using a peristaltic pump (Manostat 72-310-000; Barnant Company, Barrington, IL) into 10 ml 1× phosphate-buffered saline (PBS). The pump was set at an optimum speed to allow one cigarette to burn in approximately 15 min. The resultant extract (10 ml) was sterile filtered through a Millipore (0.22-μm filter) sterile tube to remove large particulate matter and bacteria. This solution was defined as 100% CSE. CSE at a 10% concentration was used immediately for cell treatments. The CSE preparation was standardized by measurement of the absorbance at 211 nm (optical density, 0.9753 ± 0.3154), using a spectrophotometer (UV 2501 PC UV-Vis instrument; Shimadzu, Columbia, MD).

Cell culture.

Human lung bronchial epithelial cells either sufficient (C38) or deficient (IB3) in CFTR activity were used (16). The IB3 cells were derived from bronchial epithelial cells from a cystic fibrosis patient and were immortalized by viral transformation. These cells possess both the ΔF508 and W1282X mutations in the CFTR gene. This cell line was also stably transfected with cDNA encoding wild-type human CFTR (C38). The cells were obtained from ATCC (by P. Zeitlin, John Hopkins University). Cells were cultured in a transwell plate, using LHC-8 medium containing 10% fetal bovine serum without antibiotics, and were maintained at 37°C with humidified air containing 5% CO2. Before treatment with 10 CFU/cell of M. pneumoniae and/or 10% CSE, the cells were replenished with fresh medium without the addition of antibiotics. Primary human small airway epithelial cells (SAEC) were purchased from Cambrex (Baltimore, MD). Cells were cultured using a basal medium (SAGM) containing bovine pituitary extract, human recombinant epidermal growth factor, retinoic acid, transferrin, insulin, epinephrine, fatty acid-free bovine serum albumin, and triiodithronine. SAEC were grown on transwell plates and treated similarly to IB3 and C38 cells with M. pneumoniae and/or CSE for 48 h.

Isolation of BALF.

Mice were sacrificed by administering 125 μl of Nembutal (50 mg/ml) intraperitoneally and were exsanguinated by cardiac puncture. Blood plasma was collected to perform urea analysis (Sigma Diagnostics, St. Louis, MO), which was used to determine the BAL dilution factor (10). BAL was performed by cannulating the trachea in situ with a 20-gauge tubing adapter. Three 1-ml aliquots of room temperature sterile PBS at a pH of 7.4 were instilled and collected by gentle aspiration. The BAL fluid (BALF) was pooled together and centrifuged at 4,000 × g for 10 min at 4°C to isolate BAL cells. An aliquot of supernatant (75 μl) was used to analyze GSH levels in the ELF, using high-performance liquid chromatography (HPLC) coupled with fluorescence detection as described below.

Serum and BALF urea concentrations.

In order to estimate actual ELF concentrations of soluble antioxidants from BALF, a dilution factor was derived from the difference between serum and BALF urea concentrations. This method is based on the assumption that urea concentrations in the vascular and ELF compartments are equivalent because of the freely diffusible nature of urea (10). A dilution factor was obtained by dividing the serum urea concentration by the BALF urea concentration. ELF concentrations were then calculated by multiplying the BALF concentration by the dilution factor. Urea concentrations in the samples were determined with a commercially available kit (Sigma Diagnostics, St. Louis, MO).

Determination of GSH and GSSG levels.

Intracellular GSH levels were determined in cell lysates. Cells were washed with PBS and sonicated in 200 μl of PBS for 30 s. Cell debris was removed by centrifugation. Cell lysate protein content was determine using a Coomassie Plus protein assay kit (Pierce, Rockford, IL). GSH levels were determined in an aliquot of apical fluid, BAL, or cell lysate (75 μl). To this aliquot, an equal volume of KPBS buffer (50 mM potassium phosphate buffer, 17.5 mM EDTA, 50 mM serine, and 50 mM boric acid at pH 7.4) was added, in addition to 10 μl of an internal standard (0.1 mM des-Gly-glutathione). The samples were then treated with 10 μl monobromobimane (3 mM in acetonitrile) and incubated in the dark for 30 min at room temperature. The derivatization reaction was stopped by the addition of 10 μl of 70% perchloric acid. The samples were then centrifuged at 16,000 × g for 10 min at 4°C to separate the protein pellet from the supernatant. The resulting supernatant was then transferred to an HPLC vial and analyzed on an HPLC instrument (model Elite LaChrom; Hitachi, San Jose, CA) coupled with a fluorometric detector (model L-2480). The samples were eluted on a Synergi 4-μm Hydro-RP 80A C18 column (150 × 4.6 mm; Phenomenex, Torrance, CA). The mobile phase A consisted of 1% acetic acid, 5% acetonitrile, and 94% water. The mobile phase B consisted of 1% acetic acid, 20% acetonitrile, and 79% water. Both mobile phases were adjusted to a pH of 4.25 by using ammonium hydroxide. An injection volume of 5 μl and a flow rate of 1.0 ml/minute were used to detect GSH levels. The detector excitation and emission wavelengths were set at 390 and 480 nm, respectively (19, 46). The apical and BAL GSSG levels were measured using a recycling assay in which the samples were pretreated with a GR system prior to their derivatization with monobromobimane (44).

Analysis of DNA oxidation in lung tissue and cells.

DNAs from mouse lung tissue and IB3 and C38 cells were extracted using a DNA WB extractor kit (Wako). The purified DNAs were then hydrolyzed to nucleosides with 4 units of nuclease P1 at 65°C for 20 min and 4 units of alkaline phosphatase at 60°C for 30 min. The samples were then analyzed for 2-deoxyguanosine (2dG) and 8-hydroxy-2-deoxyguanosine (8OH2dG) by HPLC coupled with UV and electrochemical detection (CoulArray model 5600; ESA Inc., Chelmford, MA), respectively. Sample analysis was performed using a 4.6- by 150-mm C18 reverse-phase column (YMCbasic; YMC Inc., Wilmington, NC). The mobile phases consisted of 0.1 M sodium acetate in 5% methanol at a pH of 5.2. 2dG was detected by UV, while 8OH2dG was detected electrochemically using an electrode potential array set at 285, 365, and 435 mV. Nucleoside concentrations were calculated using a five-point standard curve. The levels of 8OHdG were expressed as a ratio to 105 2dG bases (47).

Real-time reverse transcription-PCR analyses of GR.

Total RNA was extracted by using a Qiagen RNeasy Mini kit following treatment of cells with CSE and/or M. pneumoniae infection for 48 h to detect mRNAs for GR and glyceraldehyde-3-phosphate dehydrogenase (GAPDH). The total RNA concentration was determined with a nanodrop (ND-1000) UV-Vis spectrophotometer (NanoDrop Technologies, Wilmington, DE), adjusted to 80 ng/μl, and added to standard TaqMan reagents (Applied Biosystems, Branchburg, NJ). To compare the effects of treatments on mRNA levels, all mRNA measurements were normalized with GAPDH mRNA levels contained in total RNA pools derived from each culture. The RNAs were transcribed to produce cDNA templates that were amplified by PCRs (high-capacity cDNA reverse transcription kit; Applied Biosystems), using a 9800 series thermal cycler (Applied Biosystems) and the following conditions: 10 min at 25°C, 2 h at 37°C, and 85°C for 5 s. Gene-specific TaqMan primers and probes were used in real-time PCRs to detect GR and GAPDH. To control for possible RNA or DNA contamination during processing, non-reverse transcriptase and nontemplate controls were run in parallel with the other samples.

Western blotting of GR.

Cells were homogenized with a protease inhibitor cocktail containing EDTA and centrifuged at 500 × g for 10 min following 48 h of treatment with M. pneumoniae and/or CSE. The pellet was resuspended in 250 μl of buffer (300 mM sucrose, 10 mM HEPES, 40 μg/ml phenylmethylsulfonyl fluoride, pH 7.5). A 12.5% polyacrylamide-sodium dodecyl sulfate gel was loaded with 17 μg of protein per well. Samples were run at 150 V for 60 min and transferred to a PVDF Plus membrane (Osmonics, Westborough, MA) at 100 V for 1 h. Membranes were blocked for 1 h at room temperature in 5% powdered milk dissolved in 1× Tris-buffered saline-Tween 20 (TBST). Rabbit polyclonal immunoglobulin G (IgG) antibody against GR (1:2,000) (Abcam Inc., Cambridge, MA) was applied for 2.5 h with gentle rocking. Secondary antibody (peroxidase-conjugated Affini-Pure goat anti-rabbit IgG; Jackson ImmunoResearch Laboratories, Inc., West Grove, PA) was diluted 1:35,000 in TBST and applied for 30 min with gentle rocking. All wash steps were performed in triplicate for 10 min in TBST. GR apoprotein was detected using ECL Plus Western blotting detection reagents (Amersham Biosciences, Buckinghamshire, United Kingdom). A PVDF Plus membrane was probed for GAPDH by use of rabbit polyclonal IgG (1:2,500) (Abcam Inc., Cambridge, MA) for 2.5 h with gentle rocking.

Measurement of GR activity in cells.

Cells were homogenized in 250 μl of buffer (50 mM potassium phosphate and 1 mM EDTA, pH 7.5) and centrifuged at 8,500 × g for 10 min at 4°C. Supernatant was retained for GR analysis. GR activities were measured spectrophotometrically (340 nm) from the rate of NADPH consumption by GR during the reduction of GSSG, using a commercially available kit (Oxis International). GR activity is expressed in mU/mg of sample protein, where a unit is defined as 1 μmol of NADPH consumed per minute (46).

Statistical analysis.

Data are expressed as means ± standard deviations. Analysis of variance (ANOVA), Tukey's multiple comparison test, Pearson correlation, and Student's t test statistical analyses were performed using Prism, version 5 (GraphPad, San Diego, CA). P values of <0.05 were considered statistically significant.

RESULTS

Mycoplasma infection disrupts ELF GSH responses in ETS-exposed mice.

Groups of mice were exposed to ETS for 6 h a day 5 days a week for 16 weeks. A subgroup of mice was also inoculated with M. pneumoniae (∼108 CFU) 2, 10, and 16 weeks after the start of ETS exposure. Mice exposed to ETS for 16 weeks had a twofold increase in their steady-state ELF GSH levels compared to those exposed to FA (Fig. (Fig.1A).1A). Mice infected with M. pneumoniae for 4 months showed a small but significant decrease in their ELF GSH levels compared to the FA group. Interestingly, the mice treated with both ETS and M. pneumoniae had the lowest levels of ELF GSH among all of the groups. We also measured GSSG in all of the groups. Mice that were exposed to ETS had similar ELF GSSG levels to those in mice that were exposed to FA (Fig. (Fig.1B).1B). However, mice that were exposed solely to M. pneumoniae infection had significantly higher levels of GSSG in their ELF than those in mice that were exposed to FA or ETS alone. Interestingly, mice exposed to both ETS and M. pneumoniae had the highest ELF levels of GSSG, and these ELF GSSG changes were inversely correlated with changes in ELF GSH levels (r2 = −0.8941; P < 0.0001).

FIG. 1.
Effects of chronic ETS and Mycoplasma pneumoniae infection on ELF GSH levels and lung oxidative stress in BALB/c mice. ELF GSH (A), GSSG (B), GSH/GSSG ratio (C), and lung DNA oxidation (8OH2dG/105 2dG) (D) were measured after 4 months of exposure to ETS ...

Effects of Mycoplasma on markers of lung oxidative stress in vivo.

Oxidative stress occurs as a result of imbalances between oxidants and antioxidants and was assessed in mice exposed to ETS and M. pneumoniae. A common method to evaluate the redox environment is to examine the ratio of GSH to GSSG. Lower ratios correspond to a more oxidative environment. We observed that the ETS-exposed mice had far less oxidative stress than all the other groups (Fig. (Fig.1C),1C), which corresponded with their higher ELF GSH levels (Fig. (Fig.1A).1A). This result illustrates the lung's large capacity to mount and maintain an adequate antioxidant adaptive response to a chronic oxidant exposure. On the other hand, mice exposed to M. pneumoniae infection, by itself or in combination with ETS, had significantly lower GSH/GSSG ratios, indicative of higher levels of oxidative stress than those in mice exposed to FA or ETS alone (Fig. (Fig.1C).1C). We also measured a lung tissue marker of DNA oxidation, 8OH2dG (Fig. (Fig.1D).1D). The ETS/M. pneumoniae group had the highest levels of lung DNA oxidation, which corresponded with the lowest GSH/GSSG ratio.

Role for CFTR in cigarette smoke-induced adaptive GSH responses in vitro.

Human lung bronchial epithelial cells that are CFTR sufficient (C38) or deficient (IB3) were exposed to different concentrations of CSE for 48 h to assess the role of CFTR in CSE-induced GSH efflux responses. CSE exposure for 48 h increased apical GSH steady-state levels in both CFTR-sufficient and -deficient cells, in a concentration-dependent manner (Fig. (Fig.2A).2A). There were significant increases in apical GSH levels at 10% CSE without cytotoxicity, as measured by the release of lactate dehydrogenase (Fig. (Fig.2B).2B). These data suggest that CFTR partly contributes to the CSE-induced GSH transport response and suggest the involvement of other transporters. For all further experiments, we used 10% CSE.

FIG. 2.
Role of CFTR in cigarette smoke-induced GSH transport. (A) CFTR-sufficient (C38) and -deficient (IB3) cells were exposed to increasing concentrations of CSE, and apical GSH levels were measured after 48 h of exposure. CFTR-deficient cells have a diminished ...

Effects of Mycoplasma and CSE on adaptive GSH responses in vitro.

Primary human SAEC and C38 and IB3 cells were grown in transwells and exposed to 10 CFU/cell M. pneumoniae and 10% CSE for 48 h. The 10% CSE caused increases in the steady-state apical GSH levels (Fig. (Fig.3A).3A). The highest responses were observed in the SAEC, and the lowest responses were observed in the CFTR-deficient IB3 cells. M. pneumoniae infection alone did not show any statistically significant difference in apical GSH levels compared to the control group for all three cell systems. Interestingly, M. pneumoniae infection in combination with CSE significantly blunted the CSE-induced increase in GSH steady-state levels in all three cell systems (Fig. (Fig.3A).3A). This effect was similar to that observed in mice exposed to a combination of M. pneumoniae and ETS for 16 weeks (Fig. (Fig.1A).1A). These changes in GSH steady-state levels corresponded to changes in the level of DNA oxidation (Fig. (Fig.3B).3B). It was interesting that M. pneumoniae infection by itself did not caused DNA oxidation in SAEC but that it did so in C38 cells.

FIG. 3.
Effects of Mycoplasma pneumoniae (Mp) infection on cigarette smoke adaptive responses in vitro. Human lung bronchial epithelial cells sufficient (C38) and deficient (IB3) in CFTR and primary SAEC were exposed to 10 CFU/cell of M. pneumoniae and/or 10% ...

Mycoplasma inhibits ETS-mediated GR response.

GSH steady-state levels are controlled by rates of GSH synthesis, recycling, utilization, transport, and degradation (38, 39). Given the large change in the ratio of GSH to GSSG seen in vivo and that GR recycles GSSG back into GSH, we examined the effects of M. pneumoniae on GR at the transcription and translation levels, as well as enzymatic activity. To better understand how M. pneumoniae could disrupt normal CSE-induced GSH adaptive responses, we also compared M. pneumoniae effects to those of a known GR inhibitor, 1,3-bis(2-chloroethyl)-1-nitrosourea (BCNU). Cells were exposed to CSE alone or in combination with the GR inhibitor. Apical and intracellular GSH levels were determined 48 h later. It was interesting that BCNU significantly blocked the CSE-induced increases in apical and intracellular GSH levels (Fig. 4A and B).

FIG. 4.
Effects of GR inhibition on cigarette smoke-induced GSH adaptive responses. CFTR-sufficient (C38) cells were treated with the GR inhibitor (100 μM BCNU) in presence or absence of 10% CSE. Apical GSH levels (A) and intracellular GSH levels ...

We were interested in understanding the mechanism(s) behind M. pneumoniae infection-induced decreases in GSH levels in the apical compartment. Using GR-specific TaqMan probes, we examined the mRNA levels in CFTR-sufficient cells. CSE caused significant increases in the mRNA level of GR (Fig. (Fig.5A).5A). M. pneumoniae infection by itself did not alter the mRNA level of GR; however, in combination with CSE, it significantly abolished the CSE-induced increases in GR mRNA levels (Fig. (Fig.5A).5A). CSE-induced increases in mRNA levels of GR were associated with increases in GR protein expression, as determined by Western blotting (Fig. (Fig.5B).5B). M. pneumoniae infection in combination with CSE significantly attenuated increases in GR protein expression (Fig. (Fig.5B).5B). We also examined the GR activity in vitro and in vivo. Human epithelial cells (SAEC and C38 cells) were exposed to 10 CFU/cell M. pneumoniae and 10% CSE. After 48 h of exposure, intracellular GR activity was measured. Interestingly, we saw increases in GR activity in CSE-exposed cells which corresponded to increases in mRNA and protein expression for GR and in apical GSH levels (Fig. (Fig.5C).5C). Furthermore, we also observed that M. pneumoniae infection significantly blunted these CSE-induced increases in intracellular GR activity (Fig. (Fig.5C).5C). To test this observation in vivo, we examined lung tissue GR activity in mice exposed to ETS for 16 weeks. We observed similar effects to those seen in the in vitro studies, as ETS-exposed mice had higher lung tissue GR activities than those in groups exposed to FA. These ETS-induced increases in GR activity were completely blocked by M. pneumoniae infection (Fig. (Fig.5D5D).

FIG. 5.
Mycoplasma pneumoniae (Mp) interferes with cigarette smoke-mediated increase in GR. CFTR-sufficient (C38) cells were treated with 10 CFU/cell of M. pneumoniae and/or 10% CSE for 48 h. mRNA levels for GR (A), densitometry of protein expression ...

DISCUSSION

These studies found that (i) the lung increases ELF GSH steady-state levels when chronically exposed to ETS, and this increase is largely independent of CFTR activity; (ii) M. pneumoniae infection synergizes with ETS and is associated with enhanced oxidative stress; and (iii) M. pneumoniae causes mild oxidative stress and also inhibits ETS-induced induction of GR activity. These studies are the first to demonstrate that the common respiratory pathogen M. pneumoniae produces lung oxidative stress which synergizes with ETS through inhibition of GR and potentiates lung oxidative stress.

Epidemiological studies have shown that approximately 80% of COPD patients are current or former smokers, but only 20% of smokers actually develop COPD (40). It remains unclear why only some smokers develop COPD. Paradoxically, we and others have found that chronic ETS actually raises antioxidant steady-state levels in the lung, which can be attributed to a robust adaptive response to chronic oxidant exposure (6). We propose that factors which interfere with the lung's ability to adapt to ETS-induced challenges can make individuals more susceptible to chronic lung diseases. The present study was undertaken to investigate how respiratory pathogens such as M. pneumoniae, which produces oxidative stress and is known to exacerbate lung disease, may interfere with the lung's ability to adapt to ETS. Previous studies have shown that COPD patients may respond differently to community-acquired infections (28, 31).

M. pneumoniae infections have been found in both asthma and COPD subjects (9, 18, 20, 27, 43). In addition, M. pneumoniae infection is known to produce hydrogen peroxide via NAD (NADH2) oxidase activity (29). We observed that M. pneumoniae infection had a mild effect by itself on ELF GSH steady-state levels. However, M. pneumoniae in combination with ETS blunted the ETS-induced GSH adaptive response and prevented the lungs from maintaining adequate scavenging of ETS oxidants. Rahman and colleagues have shown that CSE can modulate the intracellular GSH level after 24 h of CSE treatment in human alveolar epithelial cells (37). We observed similar results for human lung epithelial cells after exposure to CSE. However, M. pneumoniae infection interfered significantly with both the intracellular and extracellular GSH steady-state levels, signifying the importance of GSH synthesis in maintaining GSH in both compartments. In fact, our studies suggest that alterations in GSH transport and recycling pathways can contribute to the loss of adaptive GSH responses to ETS.

A number of studies have shown that high ELF GSH concentrations similar to those found in smokers protect cultured epithelial cells against oxidative stress (6, 33, 34). We found that mice exposed to 16 weeks of ETS had increased ELF GSH steady-state levels that were blocked in the M. pneumoniae group. ETS has been shown to acutely depress ELF GSH levels, which return or rebound within 2 to 6 h (3, 12, 23). These changes have also been shown to correlate with the elevation of transcription factors known to regulate genes involved in GSH synthesis (51). In human alveolar epithelial cells, CSE exposure for 24 h caused an increase in GSH levels due to increases in γ-glutamate-cysteine ligase activity (37). In the present study, we also observed that CSE increased GSH levels both in vivo and in vitro. ELF is the first point of contact between the lung and oxidants derived from cigarette smoke, and it is not surprising to see that the lungs are able to maintain high levels of antioxidants in the ELF as part of an adaptive mechanism. Cantin and coworkers recently reported that exposure to cigarette smoke decreased CFTR mRNA expression in calu-3 cells and that this was associated with increases in cellular GSH levels, but they did not measure the extracellular GSH levels (4, 5). It is interesting that in our in vitro studies, the CFTR-deficient (IB3) cells also responded to CSE by increasing the extracellular GSH level, suggesting that there is another unidentified GSH transporter(s) responsible for the efflux of GSH, thus allowing the lung to maintain higher steady-state levels of GSH in the apical compartment (25, 47).

The GSH/GSSG ratio is routinely used as a marker of oxidative stress (15, 24). Mice exposed to M. pneumoniae infection alone or in combination with ETS were under severe oxidative stress, as evidenced by their low ELF GSH/GSSG ratios. The presence of oxidative stress in vivo and in vitro was also confirmed by increased levels of lung tissue DNA oxidation. It was interesting that mice who received both ETS and M. pneumoniae had significantly higher levels of lung DNA oxidation. A similar trend was also observed in human lung epithelial cells, and this corresponded with lower levels of apical GSH. M. pneumoniae may disrupt the lung's ability to maintain and modulate ELF GSH steady-state levels, resulting in an imbalance between oxidants and antioxidants, facilitating lung oxidative stress by producing reactive oxygen species such as hydrogen peroxide. However, it has been shown that ETS alone can also cause DNA oxidation in the tissues of mice and rats (22, 35). A possible explanation for these contradictory results may be due to the fact that Howard and colleagues used a single short-term ETS exposure. In fact, it is well known that immediately following ETS exposure, there is a sudden drop in tissue GSH levels (7). Hence, it is not surprising to see that in their experimental model, ETS-exposed mice had high levels of DNA oxidation. However, in the long-term multiple exposure system, it is possible that over time the lung can actually adapt itself against the oxidant burden by increasing steady-state GSH levels and repair systems and thus can limit oxidative stress.

When human lung epithelial cells were exposed to 10% CSE in the presence or absence of a GR enzyme inhibitor, it was evident that not only did the inhibitor interfere with the CSE-induced increases in intracellular GSH levels, but it also decreased the extracellular GSH levels. This novel finding illustrates the importance of this pathway in maintaining GSH adaptive responses to environmental oxidants such as ETS. Almagor and colleagues have shown that M. pneumoniae infections significantly decrease the host cell's catalase activity and increase hydrogen peroxide levels, resulting in oxidative damage (2). They also demonstrated that this was followed by an increase in extracellular GSSG level and a decreased GSH/GSSG ratio, but the possible mechanism(s) was not investigated (1). Our data are in agreement with their studies, since we also observed increases in ELF GSSG levels in M. pneumoniae-infected mice and in ETS-treated and M. pneumoniae-infected mice which were associated with decreases in the GSH/GSSG ratio and increases in DNA oxidation. Many transcription factors require a proper redox state to function (30). It may be possible that M. pneumoniae interferes with the ETS adaptive response through this type of mechanism. We observed that M. pneumoniae infections alone did not cause any changes in intracellular or extracellular GR activity. These data are in agreement with a previous study published by Almagor and colleagues where they demonstrated with human fibroblasts that M. pneumoniae infections did not cause changes in the GR activity (1). Recently, Singh and coworkers showed that ETS can induce GPX-2 in the lungs via the NF-E2-related factor (Nrf2) pathway. They also showed that Nrf2+/+ mice were more resistant to ETS-induced oxidative stress than Nrf2−/− animals were, which was attributed to increases in GPX-2 expression as a part of the lung's adaptive response (41). We also observed that CSE caused increases in the GR activity and that M. pneumoniae blocked this adaptive response. Thus, it is interesting to speculate that M. pneumoniae may abrogate the lung's adaptive response at the transcriptional level.

It has been hypothesized that environmental factors play an important role in predisposing smokers to the development of progressive lung diseases. Atypical bacterial infections, such as Chlamydia pneumoniae and M. pneumoniae, have been implicated in the exacerbation of progressive lung diseases (9, 49). For the first time, we have shown that an environmental pathogen, Mycoplasma pneumoniae, can interfere with the lung adaptive response to cigarette smoke-derived oxidants. We have also shown that M. pneumoniae infection interferes with GR activation, an important GSH adaptive enzyme, and prevents the lung from maintaining steady-state GSH levels, leading to oxidative stress (Fig. (Fig.6).6). The results presented here show that atypical bacterial infection can modulate the ELF GSH levels and enhance oxidative stress. Future medical therapies aimed at clearing bacterial infection in smokers and/or restoring ELF GSH levels may help in minimizing the oxidative stress and preventing the progression of chronic lung diseases.

FIG. 6.
GSH adaptive response to cigarette smoking and effects of Mycoplasma pneumoniae (Mp) infection. As part of the adaptive response, the lungs try to maintain the redox environment by mounting and maintaining high levels of GSH and GR. M. pneumoniae infection ...

Acknowledgments

This work was supported in part by NIH grants HL075523 (B.J.D.), HL084469 (B.J.D.), and HL073907 (R.J.M.).

Notes

Editor: B. A. McCormick

Footnotes

Published ahead of print on 21 July 2008.

REFERENCES

1. Almagor, M., I. Kahane, C. Gilon, and S. Yatziv. 1986. Protective effects of the glutathione redox cycle and vitamin E on cultured fibroblasts infected by Mycoplasma pneumoniae. Infect. Immun. 52240-244. [PMC free article] [PubMed]
2. Almagor, M., S. Yatziv, and I. Kahane. 1983. Inhibition of host cell catalase by Mycoplasma pneumoniae: a possible mechanism for cell injury. Infect. Immun. 41251-256. [PMC free article] [PubMed]
3. Bilimoria, M. H., and D. J. Ecobichon. 1992. Protective antioxidant mechanisms in rat and guinea pig tissues challenged by acute exposure to cigarette smoke. Toxicology 72131-144. [PubMed]
4. Cantin, A. M., G. Bilodeau, C. Ouellet, J. Liao, and J. W. Hanrahan. 2006. Oxidant stress suppresses CFTR expression. Am. J. Physiol. Cell Physiol. 290C262-C270. [PubMed]
5. Cantin, A. M., J. W. Hanrahan, G. Bilodeau, L. Ellis, A. Dupuis, J. Liao, J. Zielenski, and P. Durie. 2006. Cystic fibrosis transmembrane conductance regulator function is suppressed in cigarette smokers. Am. J. Respir. Crit. Care Med. 1731139-1144. [PubMed]
6. Cantin, A. M., S. L. North, R. C. Hubbard, and R. G. Crystal. 1987. Normal alveolar epithelial lining fluid contains high levels of glutathione. J. Appl. Physiol. 63152-157. [PubMed]
7. Carnevali, S., S. Petruzzelli, B. Longoni, R. Vanacore, R. Barale, M. Cipollini, F. Scatena, P. Paggiaro, A. Celi, and C. Giuntini. 2003. Cigarette smoke extract induces oxidative stress and apoptosis in human lung fibroblasts. Am. J. Physiol. Lung Cell Mol. Physiol. 284L955-L963. [PubMed]
8. Cassell, G. H., and B. C. Cole. 1981. Mycoplasmas as agents of human disease. N. Engl. J. Med. 30480-89. [PubMed]
9. Cherry, J. D., D. Taylor-Robinson, H. Willers, and A. C. Stenhouse. 1971. A search for mycoplasma infections in patients with chronic bronchitis. Thorax 2662-67. [PMC free article] [PubMed]
10. Chinard, F. P. 1992. Quantitative assessment of epithelial lining fluid in the lung. Am. J. Physiol. 263L617-L618. [PubMed]
11. Chu, H. W., R. Breed, J. G. Rino, R. J. Harbeck, M. R. Sills, and R. J. Martin. 2006. Repeated respiratory Mycoplasma pneumoniae infections in mice: effect of host genetic background. Microbes Infect. 81764-1772. [PubMed]
12. Cotgreave, I. A., U. Johansson, P. Moldeus, and R. Brattsand. 1987. The effect of acute cigarette smoke inhalation on pulmonary and systemic cysteine and glutathione redox states in the rat. Toxicology 45203-212. [PubMed]
13. Crystal, R. G. 1991. Oxidants and respiratory tract epithelial injury: pathogenesis and strategies for therapeutic intervention. Am. J. Med. 9139S-44S. [PubMed]
14. Daxboeck, F., R. Krause, and C. Wenisch. 2003. Laboratory diagnosis of Mycoplasma pneumoniae infection. Clin. Microbiol. Infect. 9263-273. [PubMed]
15. Day, B. J., A. M. van Heeckeren, E. Min, and L. W. Velsor. 2004. Role for cystic fibrosis transmembrane conductance regulator protein in a glutathione response to bronchopulmonary Pseudomonas infection. Infect. Immun. 722045-2051. [PMC free article] [PubMed]
16. Flotte, T. R., S. A. Afione, R. Solow, M. L. Drumm, D. Markakis, W. B. Guggino, P. L. Zeitlin, and B. J. Carter. 1993. Expression of the cystic fibrosis transmembrane conductance regulator from a novel adeno-associated virus promoter. J. Biol. Chem. 2683781-3790. [PubMed]
17. Folkerts, G., J. Kloek, R. B. Muijsers, and F. P. Nijkamp. 2001. Reactive nitrogen and oxygen species in airway inflammation. Eur. J. Pharmacol. 429251-262. [PubMed]
18. Gil, J. C., R. L. Cedillo, B. G. Mayagoitia, and M. D. Paz. 1993. Isolation of Mycoplasma pneumoniae from asthmatic patients. Ann. Allergy 7023-25. [PubMed]
19. Gupta, S., L. K. Rogers, S. K. Taylor, and C. V. Smith. 1997. Inhibition of carbamyl phosphate synthetase-I and glutamine synthetase by hepatotoxic doses of acetaminophen in mice. Toxicol. Appl. Pharmacol. 146317-327. [PubMed]
20. Hahn, D. L. 1998. Chlamydia pneumoniae and asthma. Thorax 531095-1096. [PMC free article] [PubMed]
21. Hickman-Davis, J. M., C. McNicholas-Bevensee, I. C. Davis, H. P. Ma, G. C. Davis, C. A. Bosworth, and S. Matalon. 2006. Reactive species mediate inhibition of alveolar type II sodium transport during mycoplasma infection. Am. J. Respir. Crit. Care Med. 173334-344. [PMC free article] [PubMed]
22. Howard, D. J., L. A. Briggs, and C. A. Pritsos. 1998. Oxidative DNA damage in mouse heart, liver, and lung tissue due to acute side-stream tobacco smoke exposure. Arch. Biochem. Biophys. 352293-297. [PubMed]
23. Ishizaki, T., Y. Kishi, F. Sasaki, S. Ameshima, T. Nakai, and S. Miyabo. 1996. Effect of probucol, an oral hypocholesterolaemic agent, on acute tobacco smoke inhalation in rats. Clin. Sci. (London) 90517-523. [PubMed]
24. Jones, D. P. 2002. Redox potential of GSH/GSSG couple: assay and biological significance. Methods Enzymol. 34893-112. [PubMed]
25. Kogan, I., M. Ramjeesingh, C. Li, J. F. Kidd, Y. Wang, E. M. Leslie, S. P. Cole, and C. E. Bear. 2003. CFTR directly mediates nucleotide-regulated glutathione flux. EMBO J. 221981-1989. [PMC free article] [PubMed]
26. Kraft, M., G. H. Cassell, J. E. Henson, H. Watson, J. Williamson, B. P. Marmion, C. A. Gaydos, and R. J. Martin. 1998. Detection of Mycoplasma pneumoniae in the airways of adults with chronic asthma. Am. J. Respir. Crit. Care Med. 158998-1001. [PubMed]
27. Lambert, H. P. 1968. Antibody to Mycoplasma pneumoniae in normal subjects and in patients with chronic bronchitis. J. Hyg. (London) 66185-189. [PMC free article] [PubMed]
28. Lieberman, D., M. Ben-Yaakov, Z. Lazarovich, S. Hoffman, B. Ohana, M. G. Friedman, B. Dvoskin, M. Leinonen, and I. Boldur. 2001. Infectious etiologies in acute exacerbation of COPD. Diagn. Microbiol. Infect. Dis. 4095-102. [PubMed]
29. Low, I. E., and S. M. Zimkus. 1973. Reduced nicotinamide adenine dinucleotide oxidase activity and H2O2 formation of Mycoplasma pneumoniae. J. Bacteriol. 116346-354. [PMC free article] [PubMed]
30. Lyakhovich, V. V., V. A. Vavilin, N. K. Zenkov, and E. B. Menshchikova. 2006. Active defense under oxidative stress. The antioxidant responsive element. Biochemistry (Moscow) 71962-974. [PubMed]
31. Meloni, F., E. Paschetto, P. Mangiarotti, M. Crepaldi, M. Morosini, A. Bulgheroni, and A. Fietta. 2004. Acute Chlamydia pneumoniae and Mycoplasma pneumoniae infections in community-acquired pneumonia and exacerbations of COPD or asthma: therapeutic considerations. J. Chemother. 1670-76. [PubMed]
32. Mogulkoc, N., S. Karakurt, B. Isalska, U. Bayindir, T. Celikel, V. Korten, and N. Colpan. 1999. Acute purulent exacerbation of chronic obstructive pulmonary disease and Chlamydia pneumoniae infection. Am. J. Respir. Crit. Care Med. 160349-353. [PubMed]
33. Morrison, D., I. Rahman, S. Lannan, and W. MacNee. 1999. Epithelial permeability, inflammation, and oxidant stress in the air spaces of smokers. Am. J. Respir. Crit. Care Med. 159473-479. [PubMed]
34. Neurohr, C., A. G. Lenz, I. Ding, H. Leuchte, T. Kolbe, and J. Behr. 2003. Glutamate-cysteine ligase modulatory subunit in BAL alveolar macrophages of healthy smokers. Eur. Respir. J. 2282-87. [PubMed]
35. Park, E. M., Y. M. Park, and Y. S. Gwak. 1998. Oxidative damage in tissues of rats exposed to cigarette smoke. Free Radic. Biol. Med. 2579-86. [PubMed]
36. Pryor, W. A., and K. Stone. 1993. Oxidants in cigarette smoke. Radicals, hydrogen peroxide, peroxynitrate, and peroxynitrite. Ann. N. Y. Acad. Sci. 68612-27. [PubMed]
37. Rahman, I., A. Bel, B. Mulier, M. F. Lawson, D. J. Harrison, W. Macnee, and C. A. Smith. 1996. Transcriptional regulation of gamma-glutamylcysteine synthetase-heavy subunit by oxidants in human alveolar epithelial cells. Biochem. Biophys. Res. Commun. 229832-837. [PubMed]
38. Rahman, I., S. K. Biswas, and A. Kode. 2006. Oxidant and antioxidant balance in the airways and airway diseases. Eur. J. Pharmacol. 533222-239. [PubMed]
39. Rahman, Q., P. Abidi, F. Afaq, D. Schiffmann, B. T. Mossman, D. W. Kamp, and M. Athar. 1999. Glutathione redox system in oxidative lung injury. Crit. Rev. Toxicol. 29543-568. [PubMed]
40. Repine, J. E., A. Bast, I. Lankhorst, et al. 1997. Oxidative stress in chronic obstructive pulmonary disease. Am. J. Respir. Crit. Care Med. 156341-357. [PubMed]
41. Singh, A., T. Rangasamy, R. K. Thimmulappa, H. Lee, W. O. Osburn, R. Brigelius-Flohe, T. W. Kensler, M. Yamamoto, and S. Biswal. 2006. Glutathione peroxidase 2, the major cigarette smoke-inducible isoform of GPX in lungs, is regulated by Nrf2. Am. J. Respir. Cell Mol. Biol. 35639-650. [PMC free article] [PubMed]
42. Snider, G. L. 1989. Chronic obstructive pulmonary disease: risk factors, pathophysiology and pathogenesis. Annu. Rev. Med. 40411-429. [PubMed]
43. Teig, N., A. Anders, C. Schmidt, C. Rieger, and S. Gatermann. 2005. Chlamydophila pneumoniae and Mycoplasma pneumoniae in respiratory specimens of children with chronic lung diseases. Thorax 60962-966. [PMC free article] [PubMed]
44. Tietze, F. 1969. Enzymic method for quantitative determination of nanogram amounts of total and oxidized glutathione: applications to mammalian blood and other tissues. Anal. Biochem. 27502-522. [PubMed]
45. van der Vliet, A., and C. E. Cross. 2000. Oxidants, nitrosants, and the lung. Am. J. Med. 109398-421. [PubMed]
46. Velsor, L. W., C. Kariya, R. Kachadourian, and B. J. Day. 2006. Mitochondrial oxidative stress in the lungs of cystic fibrosis transmembrane conductance regulator protein mutant mice. Am. J. Respir. Cell Mol. Biol. 35579-586. [PMC free article] [PubMed]
47. Velsor, L. W., A. van Heeckeren, and B. J. Day. 2001. Antioxidant imbalance in the lungs of cystic fibrosis transmembrane conductance regulator protein mutant mice. Am. J. Physiol. Lung Cell Mol. Physiol. 281L31-L38. [PubMed]
48. Vineis, P., M. Alavanja, and S. Garte. 2004. Dose-response relationship in tobacco-related cancers of bladder and lung: a biochemical interpretation. Int. J. Cancer 1082-7. [PubMed]
49. Von Hertzen, L., H. Alakarppa, R. Koskinen, K. Liippo, H. M. Surcel, M. Leinonen, and P. Saikku. 1997. Chlamydia pneumoniae infection in patients with chronic obstructive pulmonary disease. Epidemiol. Infect. 118155-164. [PMC free article] [PubMed]
50. Witschi, H., I. Espiritu, R. R. Maronpot, K. E. Pinkerton, and A. D. Jones. 1997. The carcinogenic potential of the gas phase of environmental tobacco smoke. Carcinogenesis 182035-2042. [PubMed]
51. Yang, S. R., A. S. Chida, M. R. Bauter, N. Shafiq, K. Seweryniak, S. B. Maggirwar, I. Kilty, and I. Rahman. 2006. Cigarette smoke induces proinflammatory cytokine release by activation of NF-kappaB and posttranslational modifications of histone deacetylase in macrophages. Am. J. Physiol. Lung Cell Mol. Physiol. 291L46-L57. [PubMed]

Articles from Infection and Immunity are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try

Formats:

Save items

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...