Department of Cell Biology, Duke University Medical Center, Durham, North Carolina 27710
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ABSTRACT |
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Pulmonary surfactant participates in the regulation of alveolar compliance and lung host defense. Surfactant homeostasis is regulated through a combination of synthesis, secretion, clearance, recycling, and degradation of surfactant components. The extracellular pool size of surfactant protein (SP) D fluctuates significantly during acute inflammation. We hypothesized that changes in SP-D levels are due, in part, to altered clearance of SP-D. Clearance pathways in rats were assessed with fluorescently labeled SP-D that was instilled into control lungs or lungs that had been treated with lipopolysaccharide (LPS) 16 h earlier. SP-D clearance from lavage into lung tissue was time dependent from 5 min to 1 h and 1.7-fold greater in LPS-treated lungs than in control lungs. Analysis of cells isolated by enzymatic digestion of lung tissue revealed differences in the SP-D-positive cell population between groups. LPS-treated lungs had 28.1-fold more SP-D-positive tissue-associated neutrophils and 193.6-fold greater SP-D association with those neutrophils compared with control lungs. These data suggest that clearance of SP-D into lung tissue is increased during inflammation and that tissue-associated neutrophils significantly contribute to this process.
type II cells; neutrophils; macrophages; surfactant recycling; surfactant protein A; lipopolysaccharide
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INTRODUCTION |
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PULMONARY SURFACTANT is a lipid, protein, and carbohydrate complex that reduces surface tension at the alveolar air-liquid interface and helps regulate pulmonary host defense. Homeostasis of the surfactant pool is mediated through synthesis, secretion, clearance, recycling, and degradation of surfactant components (12, 21, 26, 32). Disruptions to alveolar surfactant pool size and composition can be initiated by a variety of mechanisms including chronic and acute disease states (13, 18, 20). In the following studies, we investigated the effects of acute inflammation on the clearance of surfactant protein (SP)-D from the extracellular pulmonary surfactant pool.
SP-D is a large oligomeric lectin that functions in both roles of pulmonary surfactant through regulation of surfactant homeostasis (1, 14) and modulation of the innate immune response in the lung (2, 31). SP-D is synthesized in alveolar type II cells and nonciliated airway epithelial cells (Clara cells) of rat lungs (17, 30) and has been localized to type II cells, Clara cells, and alveolar macrophages by immunocytochemistry (3, 29). In vitro studies have demonstrated binding, internalization, and degradation of SP-D by macrophages (6, 15) and type II cells, which also recycle SP-D to surfactant storage and secretion organelles termed lamellar bodies (9).
Lipopolysaccharide (LPS)-induced acute pulmonary inflammation causes rapid changes in the composition of the surfactant pool and the resident cell population of the lung. Alterations in the levels of surfactant components during inflammation are quite complex because surfactant lipids, SP-A, and SP-D appear to be regulated through independent mechanisms (20, 27, 28). McIntosh and coworkers (20) reported a significant drop in the amount of SP-D recovered from bronchoalveolar lavage fluid 6 h after LPS treatment, a return to normal levels within 24 h, and a dramatic rise above normal 72 h after treatment. Whether these changes are regulated through changes in secretion, clearance, recycling, degradation, or a combination of mechanisms is not known. Inflammation-induced changes in lung cell populations could also influence the homeostasis of the surfactant pool. Lavage-associated cells from normal lungs are predominantly macrophages, whereas LPS treatment induces the migration of neutrophils into the alveolar spaces (22) and the proliferation of type II cells (23).
In this study, we focused on the clearance of SP-D during acute inflammation in an LPS-treated rat lung model and used fluorescently labeled SP-D to track metabolism of the SP-D pool. We hypothesized that fluctuations in SP-D pool size during inflammation are due, in part, to altered clearance of SP-D from the extracellular surfactant pool. Our results indicate that LPS-induced inflammation increases the clearance rate of SP-D by the lung and that neutrophils contribute significantly to this process.
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METHODS |
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Materials. Elastase for type II cell isolations was purchased from Worthington Biochemicals (Freehold, NJ). Dulbecco's PBS, DMEM, and fetal bovine serum (FBS) were obtained from Life Technologies (Gaithersburg, MD). Cell culture medium for the production of recombinant SP-D was from Irvine Scientific (Santa Ana, CA). Syringes, needles, tissue strainers, and dispase were purchased from Becton Dickinson (Franklin Lakes, NJ). Dispase was obtained from Collaborative Biochemical Products (Becton, MA). Low-endotoxin BSA, Percoll, thyroglobulin, 026:B6 Escherichia coli LPS, and all other chemicals were obtained from Sigma (St. Louis, MO).
SP isolation. Wild-type rat SP-D was isolated from the bronchoalveolar lavage fluid of silica-treated rats as previously described (33). Recombinant rat SP-D was expressed, purified, and characterized as previously described (6). Briefly, recombinant SP-D was purified by maltose-affinity chromatography from serum-free medium incubated with Chinese hamster ovary cells expressing a full-length rat SP-D cDNA clone (a generous gift from Dr. James H. Fisher, Denver Health Medical Center, Denver, CO). Functionality of the SP-D was assessed by its ability to aggregate Y1088 E. coli bacteria (16) (data not shown). Human SP-A was purified from the bronchoalveolar lavage fluid of patients with alveolar proteinosis as previously described (19). Functionality of SP-A was assessed by its ability to aggregate surfactant-like lipids (8) (data not shown). Purity of SP-A and SP-D preparations was assessed with Coomassie staining of SDS-PAGE gels. Representative preparations of SP-A and SP-D were analyzed for bacterial endotoxin contamination with a BioWhittaker (Walkersville, MD) Limulus amebocyte lysate assay. Endotoxin levels for all proteins measured <1 pg endotoxin/µg protein.
Alexa 546 and Alexa 488 labeling of proteins. Rat wild-type SP-D, rat recombinant SP-D, and thyroglobulin were labeled with fluorescent dyes with an Alexa Fluor 546 or Alexa Fluor 488 protein-labeling kit (Molecular Probes, Eugene, OR). Alexa dyes were selected because their magnitude of fluorescence is constant from pH 4 to 10 and they have a low level of photobleaching compared with similar fluorescent dyes used for protein labeling. Proteins were dialyzed against PBS (pH 7.4) before and again after labeling to remove unbound Alexa label. Human SP-A was labeled as indicated for the other proteins except it was dialyzed against HEPES-buffered water (5 mM, pH 7.0) before Alexa labeling and against Tris-buffered water (5 mM, pH 7.4) after labeling. Additionally, to protect against overlabeling of SP-A, the pH of the SP-A solution was not raised with sodium bicarbonate as recommended in the manufacturer's protocol. All proteins had a labeling efficiency of 5-10 Alexa molecules/protein monomer. Functionality of the labeled SP-A and SP-D was tested as described for unlabeled proteins in SP isolation, and their ability to bind freshly isolated rat alveolar macrophages in a time- and calcium-dependent manner was also tested (6, 15) (data not shown).
Intratracheal instillation of rat lungs. Male pathogen-free Sprague-Dawley rats (175-200 g; Taconic Farms, Germantown, NY) were anesthetized with halothane deeply enough that they remained unconscious throughout the procedure and lacked a cough reflex on instillation. Endotoxin (0.1 mg/kg of 026:B6 E. coli), fluorescently labeled SP-D (2 or 5 µg), SP-A (2 µg), or thyroglobulin (2 µg) was suspended in 200 µl of sterile 0.15 M saline. The total instilled dose never exceeded 250 µl of fluid, and vehicle-only controls were instilled with identical dose volumes. Instillations were conducted through an unoccluded trachea on an inclined surface and were followed by three 10-ml volumes of air to facilitate the distribution of the instilled fluid.
Analysis of lung compartments for distribution of fluorescent proteins. After instillation of fluorescently labeled proteins (2 µg), the rats were anesthetized with a lethal injection of 40 mg of pentobarbital sodium (Abbott Laboratories, North Chicago, IL) that included 1,000 U of heparin. After loss of the pinch reflex, the trachea was cannulated, the rat was exsanguinated via the renal vein, and its chest cavity was opened. The lungs were perfused through the pulmonary artery with 50-75 ml of a calcium buffer (140 mM NaCl, 5 mM KCl, 2.5 mM Na2HPO4, 10 mM HEPES, 2.0 mM CaCl2, and 1.3 mM MgSO4 at 37°C). The lungs were lavaged with three 10-ml volumes of EGTA buffer (140 mM NaCl, 5 mM KCl, 2.5 mM Na2HPO4, 10 mM HEPES, and 0.2 mM EGTA at 37°C), with each volume being recycled three times. The lavage fluid was immediately stored on ice. The lungs were removed from the chest cavity, individual lobes were dissected from the major airways, and the tissue was rinsed with ice-cold 150 mM NaCl. Lung tissue was cut into 5-mm pieces and stored in 5 ml of ice-cold lysis buffer (50 mM sodium phosphate buffer, 150 mM NaCl, 2 mM EDTA, and 0.5% Nonidet P-40, pH 7.2) until homogenized with five passes of a Wheaton Teflon pestle tissue grinder at 100 rpm. All homogenate volumes were adjusted to 8 ml with lysis buffer. Lavage fluid and lung homogenate were centrifuged at 250 g for 10 min at 4°C. The lavage fluid supernatant was collected and is henceforth referred to as "lavage fluid." The lavage fluid pellet, consisting primarily of lavage-associated cells, was collected in lysis buffer at 5 × 106 cells/ml and is referred to as "lavage cells." Lung homogenate supernatant was collected and is referred to as "lung tissue." To decrease background fluorescence, lung homogenate samples were centrifuged at 10,000 g for 15 min at 4°C, and the supernatant was collected. With the use of control samples containing known amounts of SP-A and SP-D, it was determined that centrifugation under these conditions did not detectably deplete the samples of fluorescently labeled protein (data not shown). Samples were read in a Fluoromax fluorometer (SPEX, Edison, NJ) or a FluoroCount plate reader (Packard Instrument, Meriden, CT) and compared with control samples isolated from lungs instilled with protein instillation vehicle only.
Isolation of lung cells by dispase digestion.
Normal and LPS-treated rat lungs instilled with Alexa-labeled SP-D (5 µg) were isolated and lavaged as described in Analysis of lung
compartments for distribution of fluorescent proteins. The lungs
were carefully dissected from the body, cleaned of all nonlung tissue
and blood, and lavaged with two 10-ml volumes of calcium buffer
(37°C). Lungs were then lavaged with 10 ml of dispase (undiluted,
37°C) and allowed to drain. The lungs were filled with 10 ml of
dispase and suspended in 5 ml of dispase in a 37°C water bath for 15 min, with the syringe left in the tracheal cannula to prevent drainage.
Lung tissue was removed from the major airways, cut into 5-mm pieces,
and returned to the dispase solution for 15 min at 37°C. Lung tissue
pieces were chopped 100 times with very sharp scissors in 5 ml of the
dispase solution along with 2 µg of DNase. The tissue suspension was
returned to the remaining dispase solution with 4 ml of FBS, and the
total volume was brought to 40 ml with calcium buffer. The suspension
was shaken vigorously for 2 min in a 37°C water bath and subsequently
strained through 100-, 40-, and 15-µm nylon mesh cell strainers.
Cells were centrifuged at 250 g for 10 min at 4°C and
resuspended in ice-cold DMEM for live cell fluorescence-activated cell
sorting (FACS). The viability of the cells isolated from normal and
LPS-treated lungs averaged >98%. SP-D-positive and -negative cells
were sorted and collected as outlined in FACS analysis, then
were stained and analyzed for purity as described in Cell
staining and purity analysis (Table 1).
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Isolation of lung tissue-associated neutrophils. Lung neutrophils were isolated from normal and LPS-treated rat lungs instilled with Alexa-labeled SP-D (5 µg). Cells were isolated from lungs with the protocol outlined in Isolation of lung cells by dispase digestion. The cell suspension from each lung was centrifuged and resuspended in 10 ml of ice-cold neutrophil isolation buffer (PBS with 10 mM Tris base and 0.5% BSA, pH 7.4) with 1% FBS and held on ice for 15 min. Biotin-labeled mouse anti-rat CD11b (0.25 µg/106 cells; PharMingen, San Diego, CA) was added, and cells were incubated for 15 min on ice. Cells were washed twice by centrifugation at 4°C with isolation buffer. Cells were resuspended in 2.5 ml of isolation buffer and incubated on ice with streptavidin MicroBeads (0.5 µl/106 cells; Miltenyi Biotec, Auburn, CA) for 20 min. Cells were washed twice by centrifugation at 4°C with isolation buffer and resuspended in degassed, room temperature isolation buffer, and antibody-labeled cells were isolated with an LS+ separation column (Miltenyi Biotec) according to the manufacturer's protocol.
Column-positive cells were applied to a Percoll step gradient to enrich the neutrophil population. Cells were resuspended in 4 ml of Percoll with a density of 1.052 g/ml. Percoll (4 ml) at densities of 1.072 and 1.085 g/ml was layered below the cell suspension, and the gradient was centrifuged at 500 g for 20 min at room temperature. Neutrophils were collected at the interface of the 1.072 and 1.085 g/ml steps. All Percoll solutions contained 0.15 M NaCl. Cells were fixed in 1% formaldehyde in PBS and analyzed by FACS for Alexa fluorescence. Using the protocol outlined in Cell staining and purity analysis, the purities of the neutrophil preparations from normal lungs averaged 92 ± 1.7% neutrophils, 3% macrophages, 4% lymphocytes, and 1% other cells (n = 8 repetitions). Purities of the neutrophil preparations from LPS-treated lungs averaged 97 ± 0.5% neutrophils, 1% macrophages, 1% lymphocytes, and 1% other cells (n = 8 repetitions). The viability of neutrophils from both groups averaged >98%.Isolation of lavage-associated neutrophils. Lavage-associated cells from LPS-treated lungs were applied to Percoll gradients identical to those used for neutrophil enrichment in Isolation of lung tissue-associated neutrophils. Neutrophils were collected from the same interface, fixed in 1% formaldehyde in PBS, and analyzed by FACS for Alexa fluorescence. Purity of the lavage-associated neutrophil preparations averaged 98% ± 0.5 neutrophils, 1% macrophages, and 1% lymphocytes according to the protocol used in Cell staining and purity analysis (n = 8 repetitions). The viability of lavage-associated cells averaged >99%.
Isolation of lavage-associated and lung tissue-associated macrophages. Macrophages were isolated from lavage fluid and lung tissue with the protocols described for neutrophil isolation except that macrophages were separated with the use of a modified Percoll step gradient for final macrophage population enrichment. Cells were resuspended in 4 ml of Percoll with a density of 1.068 g/ml and were layered below 4 ml of 1.039 g/ml of Percoll. The gradient was centrifuged at 500 g for 20 min at room temperature. Macrophages were collected at the interface of the 1.039 and 1.068 g/ml steps. All Percoll solutions contained 0.15 M NaCl. Cells were fixed in 1% formaldehyde in PBS and analyzed by FACS for Alexa 546 fluorescence. With the protocol outlined in Cell staining and purity analysis, the purities of the lavage-associated macrophage preparations averaged 98 ± 0.4% macrophages and 2% neutrophils (n = 8 repetitions) for normal lungs and 95 ± 1.1% macrophages, 4% neutrophils, and 1% lymphocytes for LPS-treated lungs (n = 4 repetitions). Purities of tissue-associated macrophages averaged 96 ± 2.1% macrophages, 2% neutrophils, and 2% lymphocytes for normal lungs (n = 4 repetitions), and 95% ± 1.4 macrophages, 4% neutrophils, and 1% lymphocytes for LPS-treated lungs (n = 4 repetitions). The viability of macrophages from all groups averaged >98%.
Isolation of alveolar type II cells. Type II cells were isolated from normal and LPS-treated lungs instilled with Alexa-labeled SP-D (5 µg) with the use of a previously described method (5) with minor modifications. Elastase was increased to 3,000 orcein units/lung, and the surface area for IgG panning was doubled. After panning, nonadherent cells were centrifuged at 250 g and resuspended in ice-cold buffer (PBS with 1% FBS and 0.1% BSA) at 5 × 106 cell/ml and held on ice for 30 min. FITC-labeled mouse anti-rat CD45 and FITC-labeled mouse anti-CD11b monoclonal antibodies (1 µg/106 cells; PharMingen) were added, and the cells were held on ice for 30 min. Cells were washed with ice-cold PBS twice by centrifugation and fixed in 1% formaldehyde in PBS. Cells were separated by FACS based on the FITC signal, and the type II cell-enriched FITC-negative cells were analyzed by FACS for Alexa fluorescence. Cells from rats given instillations of SP-D vehicle only were used as FACS baseline controls. Purity of the FITC-negative population from normal lungs averaged 98 ± 0.6% type II cells, 1% macrophages, and 2% lymphocytes with the protocol outlined in Cell staining and purity analysis (n = 4 repetitions). Purity of the FITC-negative cells from LPS-treated lungs averaged 98 ± 1.3% type II cells and 2% lymphocytes (n = 4 repetitions). The viability of type II cells from both groups averaged 97%.
Degradation assays. Macrophages and neutrophils were isolated from lungs that were not instilled with fluorescently labeled proteins. The cells were suspended at 5 × 106 cells/ml in degradation buffer (PBS containing 0.9 mM CaCl2, 0.5 mM MgCl2, and 0.1% BSA) and then incubated with 1 µg/ml of Alexa 488 rat recombinant SP-D for 2 h at 37°C. After incubation, the cells were separated from the medium by centrifugation (200 g for 7 min). Medium was collected and held on ice. Cells were washed three times with ice-cold degradation buffer and transferred to a new tube. Cells were resuspended in fresh degradation buffer, and cell and medium samples were precipitated with trichloroacetic acid (TCA; 10% final volume) for 30 min on ice. Samples were centrifuged at 10,000 g for 10 min. The supernatant was collected in a new tube, and the pellet was resuspended in 50 mM Tris (pH 6.8) containing 2% lauryl sulfate by heating at 60°C for 15 min with frequent vortexing of the samples. Samples were read in a FluoroCount fluorescence microplate reader (Packard Instrument, Meriden, CT) and were compared with control samples containing no SP-D. The Alexa 488 SP-D stock contained 0.4% TCA-soluble fluorescence, and this value was not altered by incubation in degradation buffer for 2 h at 37°C in the absence of cells.
FACS analysis. All FACS samples were analyzed at the Duke University Medical Center (Durham, NC) Flow Cytometry Laboratory. FITC was excited at 488 nm and analyzed at 514 nm. Alexa 546 was excited at 514 nm and analyzed at 560 nm. For live cell sorts, 1.5 × 106 SP-D-positive and -negative cells were sorted into separate tubes containing 5 ml of DMEM and were subsequently used for cell staining and purity analysis. For fixed cell assessments, 10,000 cells/sample were analyzed for relative Alexa fluorescence.
Cell staining and purity analysis. Cell staining was used to identify cell types in the lung cell preparations. A HARLECO Hemacolor kit (EM Science, Gibbstown, NJ) was used to identify macrophages, neutrophils, and lymphocytes. Type II cells were identified with a previously described Papanicolaou staining method (4). Cells were examined at ×1,000 to assess purity, counting ~200 cells/sample in randomly selected fields.
Confocal microscopy. Macrophages and neutrophils were isolated from Alexa 546 SP-D-instilled lungs as described in Isolation of lavage-associated and lung tissue-associated macrophages and Isolation of lung tissue-associated neutrophils. Cells were fixed in suspension with 4% paraformaldehyde (freshly made in PBS) for 30 min on ice. Cells were washed twice with PBS, adhered to microscope slides, and mounted in a solution of 50% glycerol in PBS. Localization of Alexa 546 SP-D in macrophages and neutrophils was evaluated with a Zeiss 410 confocal microscope with a ×100 objective.
Statistics. All data reported are means ± SD. Repetitions (n) used to calculate means ± SD were from independent experiments, not from duplicates within an experiment. Comparison of a single experimental group to its corresponding control was evaluated with Student's t-test. Significance was accepted at P = 0.05. Comparison of experimental groups to their corresponding controls and to each other was evaluated for differences with an analysis of variance (ANOVA). When differences were indicated by ANOVA, experimental groups were tested against the control and each other with Tukey's test of pairwise comparisons. Significance was accepted when P < 0.05.
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RESULTS |
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LPS instillation induced physical changes in rat lungs. Compared with untreated lungs, LPS-instilled lungs displayed light golden-colored areas after perfusion. Although spotting was noticeable in all lobes of the lung, its distribution was heterogeneous within each lobe as well as between lobes. Lavage fluid from LPS-treated lungs was noticeably foamier compared with lavage fluid from normal lungs.
LPS-induced acute lung inflammation increases SP-D association with
lung tissue.
Rat lungs instilled with LPS 16 h before Alexa 546-labeled
SP-D (Alexa SP-D) instillation showed enhanced association of Alexa label with lung tissue after 1 h compared with that in normal lungs and lungs instilled with LPS vehicle (Fig.
1). Both control groups had ~35% of
the recovered label associated with lung tissue and ~60% in the
lavage fluid fraction. In LPS-treated lungs, ~55% of recovered label
was found in tissue and 35% in lavage fluid. Localization of Alexa
label with lavage-associated cells was not altered by LPS treatment. On
average, Alexa label corresponding to ~75% of the 2-µg Alexa SP-D
dose was recovered from each lung. The percent recovery did not differ
among treatment groups (data not shown).
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LPS-induced acute inflammation does not alter SP-A or thyroglobulin
association with lung tissue.
Normal and LPS-treated rats instilled with Alexa-labeled SP-A showed no
difference in lung tissue association of Alexa label between groups
after 1 h (Fig. 2). Similar results
were observed with Alexa-labeled thyroglobulin (Fig.
3), a protein of approximately the same
molecular weight as SP-A and SP-D.
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LPS-induced acute inflammation increases the number of cells recovered from lung tissue digests. Dispase digestion of lung tissue was used to examine different cell types for their role in the clearance of SP-D into lung tissue. The population of cells released during digestion of normal and LPS-treated rat lungs consisted primarily of macrophages, type II cells, neutrophils, and lymphocytes (Table 1). Total cell recovery from LPS-treated lungs was 2.1-fold higher than recovery from control lungs, with much of the difference being accounted for by an increase in neutrophils (525% of control value) and lymphocytes (192% of control value) in the LPS-treated lung population.
Approximately 10-15% of each cell population described in Table 1 was damaged during FACS before being stained for cell purity analysis. These cells could not be confidently identified and are included under "Other cells" in Table 1.LPS-induced acute inflammation increases the number of SP-D-positive cells recovered from lung tissue. The population of cells isolated by dispase digestion of normal and LPS-treated lungs was examined by FACS for incorporation of Alexa signal (Table 1). Alexa-positive cells from digests of normal lungs consisted primarily of macrophages and type II cells (Table 1). The number of Alexa-positive cells recovered from LPS-treated lungs was ~1.7-fold higher than that from normal lungs. An increase in positive neutrophils from 0.1 × 106 to 15.9 × 106 accounted for the bulk of this increase, whereas the numbers of positive type II cells and macrophages were unchanged. Lymphocytes did not appear to be extensively involved in SP-D clearance in normal or LPS-treated lungs.
FACS analysis of the relative fluorescence units (RFUs) per SP-D-positive cell for the cells shown in Table 1 indicated a significantly higher association of Alexa label with the population of cells isolated from LPS-treated lungs (data not shown). Cells from LPS-treated lungs had a 1.2-fold increase in the average number of RFUs per cell compared with cells from normal lungs. Additionally, there was a 2.1-fold increase in the total RFUs associated with cells isolated from LPS-treated lungs compared with cells isolated from normal lungs (LPS-treated was different from normal, P = 0.03).LPS-induced acute inflammation enhances SP-D association with
tissue-associated neutrophils.
Neutrophils were examined by FACS as a possible mediator of enhanced
SP-D clearance during acute inflammation (Fig.
4). Neutrophils isolated from the tissue
of LPS-treated lungs showed an increase in the percentage of
Alexa-positive cells as well as an increase in the average number of
RFUs per cell compared with neutrophils isolated from normal lung
tissue. Taking into account the number of neutrophils isolated from
normal and LPS-treated lungs by dispase digestion (Table 1), there was
a 194-fold increase in the amount of label associated with isolated
neutrophils in response to LPS treatment.
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SP-D clearance differs in tissue-associated and lavage-associated neutrophils. Neutrophils were isolated from the lavage fluid and lung tissue of the same rat and were analyzed by FACS for association of Alexa label (Fig. 4). Of neutrophils isolated from lavage fluid, 55% were positive for the Alexa label, whereas only 16% of tissue-derived neutrophils were Alexa positive. However, the tissue-derived neutrophils had a 6.6-fold increase in number of RFUs per cell compared with neutrophils isolated from lavage fluid in the same lung.
LPS-induced acute inflammation decreases SP-D association with type
II cells.
Type II cells were examined by FACS as a possible route of enhanced
SP-D clearance into lung tissue during acute inflammation (Table
2). LPS-treated lungs had a lower
percentage of Alexa-positive type II cells than normal lungs, and the
number of RFUs per cell was 2.7-fold lower than that in type II cells
from normal lungs.
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LPS-induced acute inflammation decreases SP-D association with
macrophages.
Macrophages isolated from the lavage fluid and lung tissue of normal
and LPS-treated rats were analyzed for clearance of SP-D (Table
3). Macrophages isolated from the lavage
fluid of LPS-treated lungs had 20% less associated fluorescence than
macrophages isolated from the lavage fluid of normal lungs. Similarly,
the fluorescence associated with macrophages isolated from the tissue
of LPS-treated lungs was 28% lower than that in macrophages isolated
from the lavage fluid of normal rat lungs.
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SP-D is internalized by macrophages and neutrophils.
Confocal microscopy was used for a qualitative assessment of SP-D
internalization by macrophages and neutrophils (Fig.
5). Examination of macrophages and
neutrophils isolated from Alexa 546 SP-D-instilled lungs revealed
accumulation of Alexa fluorescence in intracellular structures.
Macrophages displayed fluorescence in numerous vesicles heterogeneous
in size and shape, whereas neutrophils displayed light vesicular
staining accompanied by a diffuse cytoplasmic staining. Little cell
surface staining was observed with either cell type. Macrophages and
neutrophils isolated by identical methods but from lungs not instilled
with Alexa 546 SP-D showed no detectable signal when examined under the
same confocal settings used for labeled cells (data not shown).
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LPS-induced acute inflammation alters uptake and degradation of
SP-D by neutrophils in vitro.
Macrophages and neutrophils were isolated from rat lungs and analyzed
for degradation of SP-D in vitro (Table
4). Cells were incubated in the presence
of Alexa 488 fluorescently labeled SP-D, and samples were analyzed for
the generation of TCA-soluble fluorescence in the media and cell
fractions as a measure of SP-D degradation. Macrophages isolated from
the lavage fluid of normal lungs degraded SP-D to a level similar to
previously reported findings (6). The percentage of
cell-associated TCA-soluble fluorescence in macrophages was 3.9-fold
higher than that in neutrophils isolated from the tissue of the same
lungs and 2.1-fold higher than the level of SP-D degradation by
neutrophils isolated from the lavage fluid of LPS-treated lungs.
However, neutrophils isolated from the tissue of LPS-treated lungs
degraded SP-D to a level similar to that of macrophages from the lavage
fluid of normal lungs.
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DISCUSSION |
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Homeostasis of the pulmonary surfactant pool is regulated by synthesis, secretion, clearance, recycling, and degradation and involves a variety of cell types. Acute inflammation disrupts the steady state of the normal lung and alters the levels of numerous surfactant components. In this study, we examined the effect of LPS treatment on the clearance of SP-D from the extracellular surfactant pool in an effort to understand further the pathways involved in the regulation of SP-D levels during inflammation.
A previous study by McIntosh and coworkers (20) reported changes in lavage fluid levels of SP-D in a rat model of LPS-induced inflammation. SP-D levels decreased 6 h post-LPS instillation, returned to normal within 24 h, and increased sixfold above normal by 72 h. Increases in SP-D mRNA levels 24 h post-LPS suggest that synthesis was increased, but the relative contributions of synthesis, secretion, and clearance were not evaluated. In this study, we characterized the clearance of SP-D in normal and acutely inflamed lungs 16 h post-LPS treatment, a time point at which immunoblot analysis showed that levels of SP-D were comparable in lavage fluid from normal and LPS-treated rats (data not shown).
It was found that intratracheally instilled Alexa-labeled SP-D was cleared from the extracellular surfactant pool more rapidly in LPS-treated lungs than in control lungs. Control experiments done with LPS vehicle alone showed that changes were not an artifact of anesthesia or instillation. SP-D localization with lung tissue was time dependent and specific to SP-D. No changes in SP-A or thyroglobulin clearance in response to LPS treatment were detected.
Analysis of cells from the enzymatic digest of LPS-treated lungs indicates that increased clearance of SP-D into the tissue is at least partially due to the association of SP-D with cells. The dispase digest released approximately the same number of macrophages and type II cells from LPS-treated lungs as normal lungs, yet LPS-treated lungs had a twofold increase in SP-D-positive cells. The increase in SP-D-positive cells from LPS-treated lungs was primarily due to increased numbers of SP-D-positive neutrophils, a cell type not extensively involved in clearance of SP-D in normal lungs.
Because of the enhanced contribution of neutrophils to the clearance of SP-D in the LPS-treated lung tissue, the finding that there was no increase in SP-D association with lavage fluid cells from LPS-treated lungs was surprising. LPS treatment induces neutrophil migration into the airspaces of the lung, increasing the number of cells recovered during lavage 10-fold. Therefore, an increase in SP-D clearance by the lavage pool of cells would be expected.
At least three factors may have contributed to our finding that association of SP-D with lavage fluid cells from LPS-treated lungs was similar to that in control lungs. First, LPS treatment did not alter the number of macrophages isolated from lavage fluid, but it did decrease SP-D association with the macrophages by ~20%. Thus the net contribution to SP-D clearance by lavage macrophages was decreased. Second, although there was a large influx of neutrophils into the airspaces, our in vitro degradation studies suggest that SP-D binding and degradation by lavage neutrophils was approximately half that of macrophages. Third, in vivo and in vitro data indicate that the ability of lavage neutrophils to internalize and degrade SP-D is much less than that of tissue-associated neutrophils isolated from the same lung. It is not clear if this difference is due to intrinsic changes in the metabolic properties of these cells or localization with different pools of SP-D in vivo. It seems likely that most lung SP-D would be localized in the alveolar space and not in a tissue compartment occupied by neutrophils. However, it is possible that the neutrophils we isolate by dispase digestion of lung tissue actually reside in the alveoli but are resistant to removal by lavage.
Type II cell clearance of SP-D was also altered during LPS-induced inflammation. We originally hypothesized that type II cells would be involved in the increased clearance of SP-D into the tissue of LPS-treated lungs because type II cells participate in the clearance and recycling of SP-D (9), but FACS analysis indicated that type II cell clearance of SP-D was decreased after LPS treatment. However, there is some conflict between our dispase and elastase digestion data as to the percentage of SP-D-positive type II cells released from normal and LPS-treated lungs. With dispase digestion, the number of SP-D-positive type II cells was similar in normal and LPS-treated lungs, whereas with elastase digestion, fewer SP-D-positive type II cells were recovered from LPS-treated lungs. Although we do not know the reason for this difference, the data suggest that clearance of SP-D by type II cells during inflammation is not increased and, in fact, may be decreased.
Using the current methods, we are not able to conclusively prove that the cell populations isolated by lung digestion are representative of cells in normal and LPS-treated lungs. However, based on results of total and SP-D-positive cells, it seems reasonable to conclude that LPS-treated lungs contain more SP-D-positive cells than normal lungs and that this is due in great part to the contribution of neutrophils to the clearance of SP-D during inflammation. Furthermore, the association of SP-D with tissue-associated neutrophils appears to be a specific interaction. LPS-treated lungs have a large increase in the total number of resident lymphocytes, yet there in no indication of SP-D clearance by lymphocytes. However, our current studies did not identify whether neutrophils and lymphocytes localized to common compartments in the lung and whether this might influence SP-D clearance by these cells.
Analysis of SP-A and thyroglobulin localization further suggests the specificity of increased SP-D clearance during inflammation. Because previous studies (7, 11, 24) have reported increased levels of SPs in the serum of patients with various pulmonary diseases, it was a concern that LPS treatment might disrupt the epithelial barrier and that SP-D would diffuse into the interstitium. SP-A and thyroglobulin were chosen to investigate inflammation-induced leakage because they are approximately the same molecular weight as SP-D. The results indicate no change in SP-A or thyroglobulin clearance into lung tissue in response to LPS treatment, suggesting that nonspecific leakage is not responsible for the change in clearance at the dose of LPS used or within the time course of the clearance study. Additionally, these data suggest that clearance of SP-A and SP-D is regulated differently during LPS-induced inflammation.
Two additional questions arise regarding the specificity of SP-D clearance by lung tissue cells. The first is whether the SP-D cleared by lung tissue-associated cells is internalized. Results of our confocal microscopy experiments suggest that SP-D is internalized by macrophages and neutrophils into intracellular structures. These data agree with a previous study by our laboratory (9) examining SP-D clearance by type II cells and macrophages. The second question is whether the Alexa 546 label might be separated from the SP-D molecule in the alveolar compartment before internalization, and thus free Alexa dye is responsible for the observed results. We examined cell-associated fluorescence using size-exclusion chromatography capable of separating Alexa 546 SP-D from Alexa 546 free dye (data not shown). The results indicated that a significant percentage of the Alexa 546 fluorescence isolated with neutrophils and macrophages corresponded to column fractions containing intact SP-D molecules. Fluorescence was also associated with fractions containing molecules smaller than intact SP-D; however, at this time, we are not certain whether this is a result of cellular degradation of SP-D, uptake of degraded SP-D components, or uptake of free Alexa dye.
Our studies of SP-A clearance revealed a slower rate of SP-A incorporation into the tissue than has been reported previously (25, 34). One possible explanation for the difference is that previous studies used ice-cold saline lavage to wash the lungs and make determinations of lavage- and tissue-associated label. In contrast, our lavage buffer contained EGTA and was instilled at 37°C. Therefore, we tested both methods to examine Alexa 546 SP-A clearance into lung tissue and found significantly higher levels of SP-A association with tissue when lungs were lavaged with cold saline compared with when lungs were lavaged with warm EGTA saline (data not shown). We hypothesize that both the increased temperature and sequestration of calcium by EGTA decreased SP-A association with lung tissue and increased the lavage fluid-associated recovery of SP-A. Additionally, lavage at 37°C may stimulate more alveolar stretch-induced surfactant secretion, specifically lamellar body exocytosis by type II cells (10).
A final point to consider is the possibility that LPS exerts a direct effect on SP-D clearance in the lung. Although it seems likely that instilled LPS had been cleared from the alveolar space after 16 h, we did not attempt to quantify the presence of LPS in our model. In addition, it is likely that some sections of the lung receive both LPS and SP-D, whereas others receive only one or, possibly, neither. We do not currently know how heterogeneous spreading of the instilled LPS and Alexa 546 doses may influence our observations or if LPS directly affects SP-D clearance.
In summary, clearance mechanisms actively contribute to the homeostasis of the extracellular pool of SP-D in the normal and acutely inflamed lung. Tissue-associated neutrophils appear to be primarily responsible for the clearance of SP-D into lung tissue, but the tissue-associated neutrophils differ greatly from lavage-associated neutrophils in their clearance and degradation of SP-D. Further studies are required to elucidate the role of other lung cells in the clearance of SP-D. Additionally, clearance of SP-D must be integrated with its synthesis, secretion, recycling, and degradation before the fluctuations in SP-D during inflammation can be fully understood.
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ACKNOWLEDGEMENTS |
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We thank Eric Walsh for excellent technical assistance in the isolation and preparation of wild-type and recombinant surfactant protein D (SP-D), Dr. Mike Cook and the staff of the Duke University Medical Center Flow Cytometry Laboratory for significant contributions toward the planning and analysis of our fluorescence-activated cell sorting experiments, and Dr. Stephen L. Young for helpful discussions and critical review of the manuscript. Additionally, we thank Dr. James H. Fisher (Denver Health Medical Center, Denver, CO) for providing the rat SP-D cDNA construct.
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FOOTNOTES |
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This work was supported by National Heart, Lung, and Blood Institute Grant R01-HL-30923.
Address for reprint requests and other correspondence: J. R. Wright, Box 3709, Dept. of Cell Biology, Duke Univ. Medical Center, Durham, NC 27710 (E-mail: j.wright{at}cellbio.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.
Received 21 September 2000; accepted in final form 14 February 2001.
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