1 Pulmonary and Critical Care Division, Department of Medicine, Duke University Medical Center and Veterans Affairs Medical Center, Durham, 27710; and 2 Pulmonary Toxicology Branch, Experimental Toxicology Division, National Health and Environmental Effects Research Laboratory, United States Environmental Protection Agency, Research Triangle Park, North Carolina 27711
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ABSTRACT |
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We investigated the role
of neutrophils in the development of endotoxin-induced airway disease
via systemic neutrophil depletion of C3H/HeBFeJ mice and coincident
inhalation challenge with lipopolysaccharide (LPS) over a 4-wk period.
Mice were made neutropenic with intraperitoneal injections of
neutrophil antiserum before and throughout the exposure period.
Experimental conditions included LPS-exposed, antiserum-treated; LPS-exposed, control serum-treated; air-exposed, antiserum-treated; and
air-exposed, control serum-treated groups. Physiological, biological,
and morphological assessments were performed after a 4-wk exposure and
again after a 4-wk recovery period. After the 4-wk exposure,
LPS-induced inflammation of the lower airways was significantly
attenuated in the neutropenic mice, although airway responsiveness (AR)
to methacholine (MCh) remained unchanged. After the recovery period,
LPS-exposed neutrophil-replete mice had increased AR to MCh when
compared with the LPS-exposed neutropenic animals. Morphometric data
indicate that the 4-wk exposure to LPS leads to a substantial expansion
of the subepithelial area of the medium-sized airways (90-129 µm
diameter) in nonneutropenic mice but not neutropenic mice, and this
difference persisted even after the recovery period. Expression of
bronchial epithelial and subepithelial transforming growth factor-1
(TGF-
1) was diminished in the challenged neutropenic mice compared
with the neutrophil-sufficient mice. These studies demonstrate that
neutrophils play a critical role in the development of chronic
LPS-induced airway disease.
lipopolysaccharide; polymorphonuclear leukocytes; endotoxin; subepithelial fibrosis; hyperreactivity; remodeling; transforming
growth factor-1
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INTRODUCTION |
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INFILTRATION OF AIRWAY MUCOSA with inflammatory cells is thought to be an important factor in the pathogenesis of a broad spectrum of airway disorders including acute and chronic bronchitis and asthma. Neutrophils [or polymorphonuclear leukocytes (PMNs)] contribute to the immune response of the airway to infectious and noninfectious irritants. Although it is usually protective and beneficial, this inflammatory response also has the potential to cause tissue injury. Neutrophils that are recruited to the sites of inflammation (airway epithelial and subepithelial regions) can cause tissue damage via the production and release of oxygen radicals, proteases, and soluble mediators of inflammation (e.g., cytokines and chemokines).
Inhaled irritants such as cigarette smoke, ozone, and bacterial endotoxin can produce inflammation through nonallergic mechanisms. Endotoxin [or lipopolysaccharide (LPS)], which is a component of the cell wall of gram-negative bacteria, is ubiquitous in the environment and is often present in high concentrations in organic dusts (40) as well as air-pollution particulate matter (2). Several studies have demonstrated that inhalation of air that is contaminated with endotoxin is associated with the classic features of asthma including reversible airflow obstruction and inflammation and persistent airway hyperreactivity and remodeling (35). Epidemiological studies have shown that the concentration of inhaled endotoxin in the bioaerosol is strongly and consistently associated with reversible airflow obstruction among cotton workers (25), agricultural workers (44), and fiberglass workers (31). Our previous studies have shown that the concentration of endotoxin in the bioaerosol is the most important occupational exposure associated with the development (43) and progression of airway disease in agricultural workers (44). Experimentally, inhalation of endotoxin can cause reversible airflow obstruction and airway inflammation in previously unexposed healthy study subjects (39). In fact, healthy study subjects challenged with dust from animal-confinement buildings develop airflow obstruction and an increase in the serum concentration of neutrophils and interleukin-6 (IL-6), all of which are most strongly associated with the concentration of endotoxin (not dust) in the bioaerosol (50). In endotoxin-sensitive (C3H/HeBFeJ) but not endotoxin-resistant (C3H/HeJ) mice, subchronic inhalation of grain dust causes persistent airway hyperreactivity and remodeling, which suggests that endotoxin is one of the principal components of grain dust that causes the development of chronic airway disease (17).
Although thickening of the subepithelial region of the airway is a consistent histological feature of asthma, cystic fibrosis, and chronic obstructive lung disease and is directly related to the clinical severity of these diseases, the biological factors that lead to a localized fibrotic response following chronic airway inflammation have not been well defined. To elucidate whether PMNs are essential to the development of chronic LPS-induced airway disease, we used PMN antiserum to produce neutrophil-depleted mice and examined the LPS-induced changes in those animals compared to similarly exposed mice that were not neutropenic. We hypothesized that antiserum to PMNs would substantially minimize the acute inflammatory response to inhaled LPS and in doing so would substantially inhibit subepithelial thickening and alter the development of chronic LPS-induced airway disease.
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MATERIALS AND METHODS |
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To determine whether neutrophils are essential to the
development of LPS-induced airway disease, we investigated the impact of systemic neutrophil depletion on chronic LPS-induced airway hyperresponsiveness, inflammation, and remodeling. Physiological, biological, and morphological measures were performed at three time
points: before the inhalation challenge, immediately after completion
of the 4-wk inhalation challenge, and 4 wk after completion of the 4-wk
inhalation challenge. Airway responsiveness (AR) to methacholine (MCh)
aerosol was determined, lung inflammation was assessed in whole lung
lavage fluid, and airway remodeling was estimated by light microscopic
morphometry, Masson-Trichrome staining, and transforming growth
factor-1 (TGF-
1) immunohistochemistry.
Animals. Forty-four male, 8-wk-old, C3H/HeBFeJ mice (a strain sensitive to endotoxin) from Jackson Laboratories (Bar Harbor, ME) were used in this study. To determine the effects of neutrophil depletion on LPS-induced airway disease, we randomly assigned the 44 mice to two experimental groups (n = 22 animals per group): one group received rabbit anti-mouse PMN antiserum, and the second group received normal rabbit serum [both sera were from Accurate Chemical and Scientific (Westbury, NY)]. In each treatment group, 16 animals were exposed to LPS and 6 were used as controls, which were exposed to filtered air. Mice in each group were evaluated before the exposure (baseline group; n = 22), immediately after the 4-wk exposure (4-wk-LPS group; n = 11), and 4 wk after the end of exposure (recovered group; n = 11). To minimize the rate of infection especially in neutropenic animals, all mice in this experiment were housed in the pathogen-free portion of the vivarium. The study protocol was in accordance with guidelines set forth by the Duke University Animal Care and Use Committee.
Generation of neutropenia. To deplete circulating neutrophils, mice assigned to the antiserum group received rabbit anti-mouse neutrophil antiserum as described previously (1, 16). Each dose was 0.5 ml of diluted antiserum (1:10 dilution in sterile saline) administered by intraperitoneal injection. The dosage and regimen of antiserum administration were chosen based on preliminary studies designed to determine optimal dosing of intraperitoneal serum to achieve functional prolonged neutropenia (>75% depletion of peripheral PMNs). Initial injections were administered 72 h before the inhalation exposure. The second dose was administered in the morning before the exposure started. The neutrophil antiserum was then administered three times a week for 4 wk for the 4-wk-LPS group of mice. The recovered group received the antiserum for an additional week (an additional three doses) after the exposure was completed. Concurrently, the control group of mice followed the same schedule and received intraperitoneal injections of 0.5 ml of diluted normal rabbit serum.
Endotoxin preparation and LPS aerosol exposures.
LPS (Escherichia coli serotype 0111:B4 from Sigma, St.
Louis, MO) was reconstituted with 10 ml of sterile PBS, and stock
aliquots (2.5 mg/ml) were stored at 20°C. Stock aliquots were
thawed once and stored at 4°C thereafter. Immediately before use, 0.7 ml of LPS stock [1.75 mg; 2,625,000 endotoxin units (EU)] were
diluted in 100 ml of PBS for nebulization. Mice were placed in
individual compartments of stainless steel wire-cage exposure racks in
two 135-l chambers: one was for LPS exposure and the other was for filtered air. Animals were exposed for 4 h per day and 5 days per
week over a 4-wk period. Endotoxin solution was aerosolized with a
constant-output atomizer (model 3076, TSI, Minneapolis, MN), and all of
the output was directed into the exposure chamber. Filtered,
dehumidified air was supplied to the nebulizer at a gauge pressure of
30 psi and a flow rate of ~3.3 l/min. The chamber was exhausted at a
flow rate of 35.0 l/min.
Endotoxin assay. The airborne concentration of endotoxin was assessed by sampling 0.3-0.4 m3 of air drawn from the exposure chamber through 25-mm binder-free glass-fiber filters held within a 25-mm polypropylene in-line air-sampling filter holder (Gelman Sciences, Ann Arbor, MI). Filters were placed in pyrogen-free petri dishes with 2 ml of sterile PBS that contained 0.05% Tween 20 (Sigma) and were then placed on a rotating shaker at room temperature for 1 h. Aliquots of the wash solution were serially diluted in pyrogen-free water and tested for endotoxin using a chromogenic Limulus amoebocyte lysate assay (QCL-1000, BioWhittaker, Walkersville, MD) according to the manufacturer's instructions. The endotoxin concentration during exposures averaged 9,477 ± 704 EU/m3 (means ± SD) over the 4-wk period, which corresponds to 6.32 ± 0.47 µg/m3 according to the Sigma certificate of analysis.
Blood collection and assessment of circulating neutrophils. Peripheral blood was first collected from all 44 mice before treatment and exposure. Additional blood samples were collected after the first couple of intraperitoneal serum (antiserum or control serum) injections before the exposure. Neutrophil concentrations were assessed once every week throughout the treatment and exposure periods. Mice were anesthetized with Metofane methoxyflurane (Mallinckrodt Veterinary, Mundelein, IL) inhalation, and ~0.2 ml of blood was drawn from the periorbital sinus of each mouse. The number of nucleated cells per cubic millimeter of blood was counted with a Bright-Line hemocytometer (Hausser Scientific, Horsham, PA). Differential counts of leukocytes were performed by counting 200 nucleated white blood cells from blood smears stained with HEMA-3 stain (Biochemical Sciences, Swedesboro, NJ).
Airway reactivity assessment.
The AR to MCh challenge was estimated in unrestrained unanesthetized
mice using whole body plethysmography. Individual mice were placed in
3-in-diameter chambers (Buxco Electronics, Troy, NY) that were
ventilated by bias airflow at a rate of 0.5 l · min1 · chamber
1. In each
plethysmograph, a pressure signal is generated from the pressure
difference between the main chamber that contains the unrestrained
mouse and a reference chamber, which cancels atmospheric disturbances.
Signals were analyzed to derive whole body respiratory parameters
(SFT3812, BioSystem XA version 2.0.2.48, Buxco), including respiratory
rate, tidal volume, inspiratory and expiratory times (TI
and TE, respectively), peak inspiratory and expiratory
flows (PIF and PEF, respectively), and relaxation time
(TR). These parameters are used by the software to
calculate enhanced pause (Penh, unitless), which strongly
correlates with lung resistance (18) and reflects changes
in pulmonary resistance during bronchoconstriction using the following
expression: Penh = [(TE
TR)/ TR] × (PEF/PIF). Lung function was
evaluated at baseline and after exposure to increasing doses of
aerosolized MCh (0, 5, 10, and 20 mg/ml). MCh was aerosolized for 1 min
and then dried for 2 min. Recording of breathing parameters began immediately after the end of MCh aerosolization and continued for 10 min. Average Penh values were determined over the first 3 min (early phase) of response to the MCh aerosol. To analyze the
reactivity of the animals to inhaled MCh, the concentration required to
induce a twofold increase (up to 200%) in control (saline-induced)
Penh was estimated in each animal by graphical interpolation from the dose-response curve as reported by others (23). This parameter was designated as the effective dose
(ED200) of MCh and was used as an index of AR. Mean ± SE values of the ED200 were determined for each
experimental group and were compared.
Whole lung lavage. Mice were euthanized by CO2 inhalation, and the trachea was exposed. The lungs were lavaged with 6.0 ml of sterile saline through a polyethylene (PE)-90 tube. The saline was infused 1 ml at a time at a pressure of 25 cmH2O. Return volume was recorded and was consistently >4.5 ml. Processing of the lavage fluid has been described previously (42). Briefly, the lavage fluid was centrifuged for 5 min at 200 g. The supernatant was decanted, and the cell pellet was resuspended with Hanks' balanced salt solution (without Ca or Mg) and washed twice. A small aliquot of resuspended cells was used for counting cells using a hemocytometer. The cell suspension (~100 µl) was spun onto a slide using a cytocentrifuge (Shanden, Southern Sewickley, PA) and was stained with HEMA-3 stain.
Tissue preparation. Lungs were perfused with saline through the pulmonary artery. Freshly prepared ice-cold 4% paraformaldehyde (Fisher Scientific, Pittsburgh, PA) in 1× PBS (pH 7.4) was instilled through the tracheal cannula into the left lung at a constant pressure of 25 cm H2O and was fixed in 4% paraformaldehyde at 4°C overnight. Tissue was embedded in paraffin, and 5-µm sections were cut and collected from the same depth in all of the tissue blocks. Sections were mounted onto positively charged slides (Superfrost Plus, Fisher Scientific, Pittsburgh, PA), stained with hematoxylin and eosin or Masson-Trichrome (IMEB, San Marcos, CA), and examined with an Optiphot-2 microscope (Nikon, Tokyo, Japan).
Morphometry: quantitative analysis.
Stereology was performed as previously described (17)
based on the earlier principals developed by Cruz-Orive and Weibel (11) and Hogg et al. (21). Airway perimeters
and wall areas were examined by capturing all conducting airway images
with a SPOT Jr. digital camera (Diagnostic Instruments, Sterling, MI) and were analyzed using Image-Pro Plus (Media Cybernetics, Silver Spring, MD) computer software. Measurements used in the study have been
previously defined (17). Briefly, internal and external airway perimeters and basement membrane perimeter were outlined and
measured at ×200 magnification. The areas of airway epithelial and
subepithelial tissue were calculated using these measurements. All
airway profiles were divided into three groups based on airway diameter: small (90 µm), medium (>90-129 µm), and large
(>129 µm). For every profile, the epithelial and subepithelial areas were normalized to the length of the adjacent basal membrane. The
length-normalized area for every airway profile as well as the mean for
each airway size was calculated for each animal. Means ± SE for
each experimental group were determined. To minimize the error that
might arise from tangential sectioning, any airway profiles that showed
a length-to-width ratio >2.5 were not used for analysis. The presence
or absence of lymphoid nodules was also taken into account and recorded
for each bronchiolar profile. Three sections, which represented
approximately the same depth of each tissue block, were stained and
analyzed by morphometry. For each mouse, 10-12 airways were
measured (mean, 10.7), and these were relatively equally distributed
between the three airway sizes (3-5 images from each airway size
contributed to the average data per animal). A total of 474 airway
profiles was obtained and measured. The epithelial and subepithelial
areas were traced and calculated over ~9 mm of the basal lamina for
each animal.
Immunohistochemical analysis.
Immunohistochemical detection of TGF-1 was performed on paraffin
sections using rabbit polyclonal affinity purified TGF-
1(V) antibody
(Santa Cruz Biotechnology, Santa Cruz, CA) and a standard avidin-biotin
protocol (Vector Laboratories, Burlingame, CA). Two sets of slides per
animal from all of the animals (2 × 44) were stained following
the protocol with the same concentrations of all the solutions and
equal incubation times for each slide. Negative-control sections were
prepared with normal rabbit IgG and were stained simultaneously.
Sections were stained with 3,3'-diaminobenzidine (DAB; Vector
Laboratories) and hydrogen peroxide and then counterstained with
hematoxylin. The patterns and intensities of TGF-
1
immunohistochemical reactions were assessed semiquantitatively using a
four-tier scoring system: 0, no reactivity; 1, weak reactivity
(light-brown staining); 2, moderate reactivity (medium-brown staining);
and 3, strong reactivity (dark-brown to black staining). All slides
were scored in a blinded fashion. The TGF-
1 signal was evaluated in
both the bronchial epithelial region and the subepithelial area, and the score was given after assessment of the entire section. A mean
value for the TGF-
1 signals was determined for each animal (from
both sets of slides), and a mean for each animal group was then calculated.
Statistical analyses. All data are expressed as means ± SE. We primarily compared the physiological and biological (inflammatory and airway morphometry) responses in PMN-depleted and control mice after a 4-wk exposure and again after a 4-wk recovery. The differences between the two variables were analyzed using the Mann-Whitney U-test (14). To analyze the data as a function of the time point after the exposure (early vs. late) and the treatment with PMN antiserum, a two-way ANOVA was performed for our outcomes of interest, which included the plethysmography data (values for ED200 of MCh), bronchoalveolar lavage (BAL) fluid cell counts and differentials, and morphometry mean values. Probability values of P < 0.05 (two-tailed) were considered statistically significant.
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RESULTS |
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Circulating blood neutrophils.
The intraperitoneal injection of rabbit anti-mouse PMN antiserum
significantly and consistently decreased the numbers of circulating neutrophils in the antiserum-treated group of mice compared with the
control serum-treated animals (Fig. 1).
Mice exposed to LPS and treated with neutrophil antiserum had
neutrophil concentrations that were 21-34% (average, 27%) of
those in control serum-treated mice. Similarly, in air-exposed mice,
neutrophils from PMN antiserum-treated mice were at circulating levels
of 24-36% (average, 31%) below the levels in air-exposed mice
treated with control serum. The total neutrophil count decreased from
3,366 ± 105 cells/mm3 in the control serum-treated
animals to 1,009 ± 27 cells/mm3 in the
antiserum-treated group (P < 0.005). Antiserum
treatment also resulted in a younger population of circulating
neutrophils; in control serum-treated mice, bands constituted
2-3% of all neutrophils, whereas in the antiserum-treated group,
bands constituted 20-24% of all neutrophils. There was no
significant difference in the number of circulating neutrophils between
LPS- and air-exposed animals injected with the same type of serum
(control serum or antiserum) at any time point. In the recovered group
of animals, the number of circulating neutrophils returned to the
baseline level 1 wk after antiserum treatment was discontinued. The
number of circulating lymphocytes in the antiserum-treated mice was
also reduced significantly (P < 0.005) to 40% (e.g.,
1,059 ± 137 cells/mm3) compared to the control
serum-treated group of animals.
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Airway physiology.
To evaluate differences in the response to inhaled MCh and provide
longitudinal comparisons, ED200 of MCh was calculated for each animal and a mean was determined for every group of mice (Fig.
2). Before exposure, there were no
significant differences in AR to aerosolized MCh between experimental
groups. AR to MCh was assessed at two time points after completion of 4 wk of LPS inhalation, within 12 h after completion of the
exposure, and at 72 h after completion of the exposure. No
consistent significant differences in airway reactivity to MCh between
antiserum- and control serum-treated animals were determined at either
the 12- or 72-h time points after completion of the inhalation
challenge. Figure 2 shows the results at the 72-h time point. However,
after the 4-wk recovery period, control serum-treated, LPS-exposed mice showed significantly increased (P < 0.005) AR to MCh
(as reflected by a decrease in ED200 of MCh) compared with
antiserum-treated, LPS-exposed mice (Fig. 2). The ANOVA comparisons
show significant (P < 0.002) time/age-dependent
changes (8 vs. 12 vs. 16 wk of age) as a decrease of the estimates of
AR to MCh within each exposure group. Further ANOVA confirmed
the univariate analyses that the only significant difference
(P < 0.001) defined from the serum treatment was found
for the LPS-exposed group after the recovery period.
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Inflammation in lower respiratory tract.
Three days after the 4-wk inhalation challenge with LPS, both control
serum-treated and antiserum-treated mice demonstrated a pronounced
pulmonary inflammatory response. LPS exposure induced a significant
increase (P < 0.05) in total BAL fluid cell number in
both control serum-treated (>25-fold) and antiserum-treated (>5-fold)
mice compared with corresponding air-exposed control animals (Fig.
3A). Changes in total BAL
fluid cell numbers in the control serum-treated, LPS-exposed group were
due predominantly to increases in the number of macrophages
(representing 60% of total BAL fluid cells), neutrophils (15%), and
lymphocytes (22%). In antiserum-treated, LPS-exposed animals, the
neutrophil influx was relatively smaller (7% of total BAL fluid
cells), whereas macrophages (75%) and lymphocytes (14%) were more
abundant. At this same time point, the LPS-exposed, antiserum-treated
mice showed a significant reduction (P < 0.005) in the
total number of BAL fluid cells and in the counts of macrophages,
neutrophils, and lymphocytes per milliliter of BAL fluid compared with
LPS-exposed animals that received control serum. Antiserum treatment
had no effect on BAL fluid cell numbers in air-exposed mice. After the completion of the 4-wk recovery period, the significant difference (P < 0.05) in total BAL fluid cell counts in
LPS-exposed vs. air-exposed animals treated with the same two types of
serum still persisted (Fig. 3B). However, the cellular
pattern was different than the response observed acutely after
exposure. Macrophages, airway epithelial cells, and lymphocytes now
comprised up to 99% of the total BAL fluid cell count, and neutrophils
contributed to <1% of the BAL fluid cell population for all treatment
groups. No significant differences in BAL fluid cell count in
antiserum-treated vs. control serum-treated animals within exposure
groups were found at the end of the recovery period. The ANOVA results
show that all univariate analyses were significant.
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Airway architecture: histopathology.
At the end of the 4-wk exposure period, control serum-treated and
antiserum-treated mice exposed to air exhibited normal lung architecture (Fig. 4A). After
LPS inhalation challenge, lungs from the nonneutropenic mice showed an
intense cellular infiltration (predominantly neutrophils) of the
interstitium and bronchiolar walls (Fig. 4B). This effect
was considerably attenuated in the neutropenic (antiserum-treated)
animals (Fig. 4C). Our morphometric analysis demonstrated
that the subepithelial area (calculated per length of basal membrane)
of the medium-size airways in the LPS-exposed, control serum-treated
group was significantly increased (P < 0.05) compared
with corresponding air-exposed control animals (Fig.
5A). Inflammatory cells and
edema fluid clearly contributed to the subepithelial area expansion,
but excluding inflammatory cells from area tracing was unavoidable. At
the same time point, mice that were LPS exposed but antiserum treated
had significantly reduced (P < 0.005) expansion of the
subepithelial area for the medium airways compared with the control
serum-treated, LPS-exposed mice.
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Immunohistological analysis.
Table 1 summarizes the average scores for
TGF-1 staining of epithelial and subepithelial compartments for all
of the experimental groups. At 72 h after a 4-wk LPS exposure,
antiserum-treated mice showed staining for TGF-
1 with moderate
intensity that was localized in the cytoplasm of the bronchiolar
epithelial cells and absent or very weak subepithelial staining (Fig.
7). At the same time point, the
LPS-exposed, control serum-treated group demonstrated weak epithelial
TGF-
1 reactivity and strong staining of the subepithelial area with
the latter significantly different (P < 0.005)
compared to the LPS-exposed, neutrophil-depleted group (Fig.
8A). After 4 wk of recovery, a
prominent TGF-
1-positive reaction in bronchiolar epithelium and an
almost lack of reactivity in the subepithelium were observed in the
sections from the lung of the LPS-exposed, control serum-treated mice.
In contrast, LPS-exposed, antiserum-treated mice showed substantially
less intense staining (P < 0.005) of the bronchiolar
epithelium compared to the neutrophil-replete, LPS-exposed animals
(Figs. 7 and 8B). As expected, staining for TGF-
1 in the
epithelial cells of bronchioli of the air-exposed animals was
considerably weaker than in the LPS-treated mice.
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DISCUSSION |
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The results of the present study indicate that the development of chronic LPS-induced airway disease is dependent on the presence of neutrophils. Mice that were depleted of circulating neutrophils had markedly less lung inflammation and demonstrated little evidence of LPS-induced airway hyperreactivity or airway remodeling (expansion of the subepithelial matrix). In contrast, neutrophil-replete mice developed airway inflammation, hyperreactivity, and remodeling after a prolonged exposure to LPS. These results provide support for the hypothesis that chronic airway disease in individuals exposed to dust contaminated with LPS may be largely mediated by PMNs and suggest a direct association between airway inflammation and remodeling.
Our study demonstrates that a 4-wk exposure to LPS by inhalation causes development of chronic airway disease in LPS-responsive mice. In the evolution of this condition, two phases could be recognized: the early or direct phase (during and immediately after the exposure), which was induced by the consistent and repetitive presence of the stimulus and during which infiltration and activation of inflammatory cells (mostly neutrophils) took place without accompanying enhancement in airway reactivity; and the late or indirect phase (4 wk after the exposure), which was characterized by fibrotic airway remodeling and hyperresponsiveness. Our results demonstrate that inhibition of neutrophil recruitment in the early phase mitigates the chronic inflammatory response to inhaled endotoxin (LPS) and subsequently attenuates the late-phase airway hyperreactivity and remodeling in LPS-responsive mice. To our knowledge, the data presented here are the first to show a temporal and causative relationship between neutrophil recruitment in the lung provoked by repetitive stimuli and the later development of airway hyperreactivity and remodeling. However, because the PMN antiserum that we used was effective but not completely selective for neutrophils, our results do not rule out the possibility that in addition to neutrophils, lymphocytes might also be involved in the development of the chronic LPS-induced airway disease.
The mechanism of action of LPS has been studied extensively. LPS is not
directly chemotactic for PMNs (10, 45); therefore, endogenous factors must be released to facilitate the extravasation of
PMNs after inhalation of LPS. Toll-like receptor 4 (TLR-4) is an
essential receptor for LPS signaling and is also the main protein
involved in recognition of gram-negative bacteria (20). Additionally, CD14, a plasma membrane protein, assists
monocyte/macrophage recognition of LPS binding protein (LBP) complexes
and enhances the response of TLR-4 to endotoxin (7).
Stimulation of TLR-4 results in activation of proinflammatory
transcriptional factors and production and release of proinflammatory
cytokines (46). In fact, in previous studies, we have
demonstrated that TNF-, IL-6, and macrophage inflammatory protein-2
(MIP-2) are released within hours of the exposure and may persist for
up to 48 h after the inhalation challenge (12). We
have also shown that recruited neutrophils are the primary cells that
produce these proinflammatory cytokines (48). The findings
presented in this report suggest that prolonged neutrophilic
inflammation of the airway wall may be an important initial step in the
development of chronic LPS-induced airway disease. This step may
initiate a chain of events that results in subsequent subepithelial
extracellular matrix deposition.
The relationship between nonallergic airway inflammation and airway hyperreactivity is poorly understood. Acute airway exposure to LPS causes airway neutrophilia and induces airflow obstruction accompanied by short-lived airway hyperreactivity to aerosolized MCh (30, 41). Although neutrophils persist 24 h after the exposure, enhancement of airway hyperreactivity to MCh was not observed (30), which suggests that neutrophils were not sufficient to cause airway hyperreactivity. This is supported by experiments that have shown that inhaled LPS, although associated with neutrophil recruitment, results in bronchial hyporesponsiveness to histamine (15). These findings suggest that the presence of neutrophils in lung compartments (air spaces, interstitium, airway and alveolar walls, and vascular bed) is insufficient to mediate increased AR to bronchoconstrictors. However, although our results demonstrate that prolonged neutrophil infiltration is also not sufficient for the development of airway hyperreactivity to MCh, the presence of neutrophils is involved in airway remodeling that eventually leads to an enhanced response to MCh. Another relevant observation in our study is the gradual decrease of airway sensitivity to inhaled MCh in mice within 8-16 wk of age. This is clearly evident in the sentinel mice included in this experiment. We are not aware of a longitudinal study on age-related changes in airway reactivity in mice to validate these findings independent of our experimental system. However, in other rodent models such as the guinea pig, the effect of age on muscarinic agonist-induced contraction of airway smooth muscle (ASM) has been demonstrated (47). In addition, in humans, it is well documented that during adolescence (6-16 yr of age) there is a natural decrease in airway reactivity to inhaled MCh (33, 34). This is true not only for normal (nonasthmatic) individuals but also for asthmatics. One of the possible mechanisms that might contribute to this phenomenon is the maturation (decrease in activity) of the Na+/K+ pump in ASM cells that occurs throughout the pubertal period (37). Those age-related changes in the function and properties of ASM cells, including changes in responsiveness to bronchoconstrictors, are reported for different species. Alternatively, this could simply reflect a change in receptor density along the airway and/or the growth of the airways.
Airway-tissue remodeling in asthma, chronic obstructive lung disease, and cystic fibrosis involves a fibrotic response that alters the structure and function of the airway. Although thickening and fibrosis of the subepithelial region beneath the basement membrane is a consistent (4, 13, 24, 29, 36, 38) histological feature of these diseases with the magnitude of this response directly related to clinical severity (6, 32), the biological factors responsible for this localized fibrotic response have not been well defined. Immunohistochemical staining reveals that the subepithelial fibrosis predominantly consists of types I, III, and IV collagen, which probably originate from fibroblasts and myofibroblasts (5, 38) rather than epithelial cells (which produce branched collagens of types V and VII; Ref. 22). In this study, we have demonstrated that the medium-size airways of nonneutropenic mice exposed to a 4-wk LPS inhalation develop, in the early phase, expansion of the subepithelial tissue area. This expansion was not observed in the neutropenic, LPS-exposed mice. Changes persisted 1 mo after the end of the exposure and were characterized by enhanced subepithelial deposition of collagen.
TGF-s are pleiotropic factors that strongly influence processes such
as somatic tissue development and repair, inflammatory diseases,
tumorogenesis, wound healing, and fibrosis (3). TGF-
1 is member of a family of several similar molecules (including TGF-
1,
-
2, and -
3) with genes that are highly conserved across species
and thought to modulate the development of fibrosis as well as other
biological processes (3). Recent studies have shown that TGF-
secreted by airway epithelia and inflammatory cells
may play a role in the development of subepithelial fibrosis. TGF-
1,
which is abundant in the lung, can be demonstrated in BAL fluid and
appears to play an important role in modulating inflammation and
fibrosis in several forms of lung disease (27, 28). The
cellular sources of TGF-
1 in the lung (especially at sites of
injury) include inflammatory cells such as lymphocytes, macrophages
(26, 49), eosinophils in the airway wall of asthmatics (32), epithelial cells (27), and platelets
(19). Our data suggest that during the early phase (the
end of the 4-wk LPS exposure) in the neutrophil-replete mice, cells in
the airway subepithelium were the predominant source of TGF-
1. This
expression of TGF-
1 in the subepithelial matrix was not observed in
the neutropenic, LPS-exposed mice. However, after 4 wk of recovery, the
bronchial epithelium of nonneutropenic mice showed strong expression of TGF-
1 whereas much less was present in the epithelium of neutropenic mice. TGF-
1 may be produced by a variety of cells that includes ASM
cells, lymphocytes, tissue macrophages, and even neutrophils (8). Once released, TGF-
1 could function in an
autocrine manner (to induce ASM cells to synthesize procollagen I) or
in a paracrine fashion as a mitogen to immature fibroblasts
(9). In aggregate, these findings suggest that expression
of TGF-
1 in the mouse conducting airways increases after chronic LPS
inhalation challenge and is dependant on neutrophilic infiltration and
associated with subsequent collagen deposition. Diminishing PMNs in the
bronchial wall with antiserum treatment reduces TGF-
1 expression and
collagen formation. We hypothesize that the increased TGF-
1 activity
induced by LPS exposure and mediated by neutrophils recruited into the tissue stimulates extracellular matrix production that contributes to
the expansion of the bronchial subepithelial region and is likely to be
critical to airway remodeling.
PMNs are phagocytic cells, and it is fully recognized that neutrophil
infiltration of the airway as a part of the acute or chronic
inflammatory response to microbial stimuli is beneficial, protective,
and important in resolving injury to the lung. Once activated, PMNs
generate free radicals and release lytic enzymes from cytosolic
granules. This ability to become activated and release damaging
intermediates is believed to mediate neutrophil-dependant host-tissue
injuries. Our findings indicate that after chronic endotoxin exposure,
neutrophils can contribute to airway remodeling and hyperreactivity.
TGF-1 may play an important role in this development. This suggests
that control of neutrophil recruitment in the airway or the
neutralization of TGF-
1 may limit the extent of chronic LPS-induced
airway disease.
![]() |
ACKNOWLEDGEMENTS |
---|
The authors thank Paul Evansky, Todd Krantz, and Najwa Haykal-Coates for excellent technical assistance in carrying out endotoxin exposures and assays.
![]() |
FOOTNOTES |
---|
This study was supported by grants from the Department of Veterans Affairs (Merit Review), the National Institute of Environmental Health Sciences (ES06537, ES07498, and ES09607), and the National Heart Lung and Blood Institute (HL-62628).
This report has been reviewed by the National Health and Environmental Effects Research Laboratory, US Environmental Protection Agency, and approved for publication. Approval does not signify that the contents necessarily reflect the views and policies of the Agency, nor does mention of trade names or commercial products constitute endorsement or recommendation for use.
Address for reprint requests and other correspondence: J. D. Savov, Duke Univ. Medical Center, PO Box 2629, Durham, NC 27710 (E-mail: jsavov{at}acpub.duke.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
10.1152/ajplung.00420.2001
Received 26 October 2001; accepted in final form 13 June 2002.
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