Departments of Medical and Surgical Sciences and Clinical Medicine and Centro per lo Studio dell' Invecchiamento, University of Padua, 35128 Padua; and Istituto di Ricovero e Cura a Carattere Scientifico, Policlinico San Matteo, University of Pavia, 27100 Pavia, Italy
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ABSTRACT |
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Alveolar macrophages degrade surfactant protein (SP) A and saturated phosphatidycholine [dipalmitoylphosphatidylcholine (DPPC)]. To clarify this process, using rabbit alveolar macrophages, we analyzed the effect of drugs known to affect phagocytosis, pinocytosis, clathrin-mediated uptake, caveolae, the cytoskeleton, lysosomal pH, protein kinase C, and phosphatidylinositol 3-kinase (PI3K) on the degradation of SP-A and DPPC. We found the following: 1) SP-A binds to the plasma membrane, is rapidly internalized, and then moves toward degradative compartments. Uptake could be clathrin mediated, whereas phagocytosis, pinocytosis, or the use of caveolae are less likely. An intact cytoskeleton and an acidic milieu are necessary for the degradation of SP-A. 2) Stimulation of protein kinase C increases the degradation of SP-A. 3) PI3K influences the degradation of SP-A by regulating both the speed of internalization and subsequent intracellular steps, but its inhibition does not prevent SP-A from reaching the lysosomal compartment. 4) The degradation of DPPC is unaffected by most of the treatments able to influence the degradation of SP-A. Thus it appears that DPPC is degraded by alveolar macrophages through mechanisms very different from those utilized for the degradation of SP-A.
surfactant protein A; dipalmitoylphosphatidylcholine; phosphatidylinositol 3-kinase
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INTRODUCTION |
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THE CONCENTRATION OF
SURFACTANT PROTEIN (SP) A in the alveoli is the net result of
secretion by type II cells and removal by type II cells and
macrophages; however, although the role of type II cells is well
established (24, 37, 41), that
of macrophages is less well understood (27). At present,
two lines of evidence indicate that macrophages might play an important role in the turnover of SP-A: first, the finding that macrophages in
vitro degrade SP-A, whereas type II cells are unable to do it
(6, 24, 40), and second, the
observation that the alveolar pool of SP-A increases in mice lacking
granulocyte-macrophage colony-stimulating factor (GM-CSF) or the common
-chain of the receptor of this growth factor (13). To
date, however, the factors that direct alveolar SP-A toward type II
cells rather than toward macrophages are unknown.
To be degraded, SP-A needs to be recognized by alveolar macrophages. Recognition could involve macrophage-specific receptors and/or receptors shared in common with type II cells. Indeed, macrophages share with type II cells a 210-kDa protein that binds the collagenous domain of SP-A in a Ca2+-dependent manner (22). In addition, they also have other SP-A receptors such as the C1q/MBL/SP-A receptor (C1qRp) that binds the collagenous domain of SP-A, surface glucoconjugates that could bind SP-A through the carbohydrate recognition domain, and organism-specific receptors associated with viruses and bacteria that could bind SP-A (27).
How macrophages degrade SP-A is still poorly known. The process is time and temperature dependent (40), seems to require an acidic milieu (because the exposure to NH4Cl is inhibitory), and is influenced by the activation state of the cell but not by the presence of surfactant lipids (6).
Besides degrading SP-A, alveolar macrophages also play a role in the degradation of surfactant lipids as indicated by the fact that they accumulate a nonmetabolizable analog of dipalmitoylphosphatidylcholine (DPPC) administered through the trachea (33) and by the finding that mice lacking GM-CSF accumulate surfactant lipids in high concentrations in the alveoli (13). Unlike type II cells that recycle a large proportion of the lipids they take up, macrophages move most of lipids they take up toward degradation (6). The degradation is time and temperature dependent (18, 29, 40), is influenced by cell energy levels, and is not as sensitive to culture conditions as the degradation of SP-A is (7). On the basis of the last observation, Bates et al. (7) suggested that once internalized by alveolar macrophages, protein and lipids could be degraded in separate compartments. Wright and Youmans (40), however, after exposing macrophages to liposomes containing fluorescent SP-A and DPPC, found that internalized labels had approximately the same cellular distribution.
In this paper, to clarify the role of macrophages in the degradation of surfactant, we exposed rabbit alveolar macrophages to substances known to affect phagocytosis, pinocytosis, clathrin-dependent and caveolae-dependent endocytosis, cytoskeletal organization, intracellular vesicular traffic, and lysosomal hydrolytic activity and then analyzed the ability to degrade SP-A and DPPC. We also studied the role of protein kinase C (PKC) that, by promoting dynamin phosphorylation, is involved in the budding of clathrin-coated vesicles (34). Finally, we studied the role of phosphatidylinositol 3-kinase (PI3K).
PI3K is a family of lipid kinases, classified as classes IA, IB, II, and III, that contains homologous catalytic domains but distinct regulatory elements and catalyzes the phosphorylation of the 3' position of the inositol ring of phosphatidylinositol or phosphoinositides (16). In Saccharomyces cerevisiae, the PI3K analog VPS-34 is involved in vesicular protein targeting (16). In mammalian cells, PI3Ks are involved in membrane ruffling (3, 21), fluid-phase pinocytosis (5, 11), phagocytosis (3), receptor-mediated endocytosis (10), intracellular traffic of vesicular organelles, (26) and signaling events that are elicited by the binding of hormones, growth factors, and cytokines to specific membrane receptors (15, 16). To study the role of PI3K, we analyzed the effect of two inhibitors, wortmannin and LY-294002, on the degradation of SP-A and DPPC by alveolar macrophages.
Our results indicate that 1) free SP-A could be taken up by macrophages by a clathrin-dependent mechanism, whereas uptake by phagocytosis or pinocytosis or through caveolae is unlikely; 2) internalized SP-A is degraded through a process that requires an intact cytoskeleton and the association with lysosomes; 3) PI3K plays an important role both during the internalization of SP-A bound to the plasma membrane and at subsequent intracellular steps, but its inhibition does not prevent SP-A from reaching the lysosomes; and 4) the degradation of DPPC is unaffected by most of the treatments able to influence the degradation of SP-A.
Thus it appears that DPPC is taken up and degraded by alveolar macrophages through mechanisms very different from those utilized for the degradation of SP-A.
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METHODS |
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Materials.
All reagents were of analytic grade. Na125I was
from Amersham Pharmacia Biotech (Little Chalfont, UK).
[3H]DPPC
{[2-palmitoyl-9,10-3H(N)]dipalmitoyl-L--phosphatidylcholine;
specific activity 40.0 Ci/mmol} was obtained from Amersham and
further purified by preparative thin-layer chromatography. Percoll was
from Pharmacia (Uppsala, Sweden). Dulbecco's modified Eagle's medium
(DMEM) was from Life Technologies (Paisley, UK). DPPC, egg
phosphatidylcholine (PC), egg phosphatidylglycerol (PG),
phosphatidylserine (PS), cholesterol (Chol), and all other chemicals
were obtained from Sigma (St. Louis, MO). Phospholipid purity was
checked by thin-layer chromatography. Lactate dehydrogenase (LDH) was
measured with a commercial kit (Roche) with pyruvate as the substrate.
Drugs.
The drugs used were 50 ng/ml of bacterial lipopolysaccharide (LPS) from
Escherichia coli 026:B6 and 100 ng/ml of interferon- (IFN-
), which stimulate phagocytosis and intracellular killing of
pathogens (1, 12, 35); 100 µM
chlorpromazine, which inhibits the endocytosis mediated by
clathrin-coated vesicles (39); 1.5 µM filipin and 1 mM
methyl-
-cyclodextrin, which inhibit caveolae-mediated endocytosis
(2); 10 µM cytochalasin D, which induces actin
depolymerization (20); 20 µM nocodazole, which inhibits
microtubule polymerization (20); 10 µM monensin, which inhibits the acidification of lysosomes (30); 100 nM
wortmannin, which irreversibly inhibits PI3K (16); 50 µM
LY-294002, a quercetin analog, which reversibly inhibits PI3K
(16, 38); and 50 ng/ml of phorbol
12-myristate 13-acetate (PMA), a stimulator of PKC (12).
The drugs were prepared as concentrated solutions and stored at
80°C. Aliquots of these compounds were thawed only once and used
immediately. Depending on the compound used, cells were exposed to
0.1% dimethyl sulfoxide or 1 µl/ml of ethanol. The drugs had
no adverse effects on viability as judged by trypan blue exclusion. LDH
release by macrophages was <0.5% of total for all treatments, with
the exception of chlorpromazine that induced the release of 10±
4% of total LDH.
Cells. Alveolar macrophages were isolated from the bronchoalveolar lavage (BAL) fluid collected from 1-kg rabbits as described by Pison et al. (31), washed with Ringer buffer albumin (RBA; 155 mM NaCl, 5 mM KCl, 1 mM MgCl2, 2 mM Na2HPO4, 10 mM glucose, 10 mM HEPES, and 1 mg bovine serum albumin/ml, pH 7.4), and used either as a suspension or after adhesion to 6- or 24-well plates. We recovered 67 × 106 ± 38 × 106 (SD) cells/kg (n = 70 samples). The cells were >90% macrophages and >95% viable.
SP-A labeling.
SP-A was isolated from the BAL fluid of patients with alveolar
proteinosis and further purified by mannose-Sepharose 4B column chromatography (4). All preparations of SP-A were tested
for LPS contamination by the Limulus amebocyte lysate assay
(Limusate, Haemachem, St. Louis, MO), and LPS concentration was always
<0.125 endotoxin unit/ml (<0.2 pg/µg protein). SP-A was labeled
with125I according to Goldstein et al. (17) to
a specific activity of 3-6 × 105 counts · min1 · µg
1. Radioactivity was
>99% precipitable with TCA and, after polyacrylamide gel
electrophoresis under reducing conditions and autoradiography, appeared
as a major band of 32-36 kDa and as a minor band of irreducible dimers weighing 64-66 kDa. 125I-SP-A was kept at 4°C
and used within 1 mo.
Association of SP-A with alveolar macrophages. Suspensions of alveolar macrophages (5 × 106 cells/ml RBA) were incubated at 37°C for 60 min in the absence and presence of 100 nM wortmannin and then exposed for 0-100 min to 1 µg 125I-SP-A/ml. After the incubation, the cells were sedimented at 500 g for 5 min, and the supernatant was saved. The cell pellet was suspended in RBA, transferred to new tubes, and washed two more times with RBA. In the supernatant, we measured the radioactivity that could not be precipitated by 20% cold TCA (TCA-soluble radioactivity). In the cells, we measured total radioactivity, TCA-soluble radioactivity, and radioactivity bound to the plasma membrane. To measure SP-A bound to the plasma membrane, 0.2-ml aliquots of the cell pellet were mixed with 4.8 ml of phosphate-buffered saline (PBS) containing 1 mg albumin/ml, pH 3.5, incubated for 5 min at 4°C, and then centrifuged at 500 g for 10 min (23). The radioactivity remaining in the supernatant represents bound SP-A released by the acid wash. Data presented are from six experiments.
Degradation of SP-A by alveolar macrophages. Alveolar macrophages (5 × 106 cells/ml RBA) were incubated at 37°C for 60 min in the absence (control) and presence of various drugs and then exposed for 60 min to 1 µg 125I-SP-A/ml. After the incubation, the cell suspensions were cooled and centrifuged at 500 g for 5 min. The resulting supernatant was recovered and used to measure TCA-soluble radioactivity. The degradation of SP-A was also studied by incubating 125I-SP-A with plain RBA (spontaneous degradation) and with medium collected from macrophages that had been incubated at 37°C for 60 min with and without drugs (conditioned medium). We routinely found that spontaneous degradation was <1% and that degradation by conditioned medium was negligible. The degradation rate, which was dependent on time and macrophage concentration, is expressed as nanograms of SP-A degraded per 106 cells per hour. The effect of various drugs is expressed as a percentage of the degradation measured in control cells. Assays were done in triplicate. Results are from 3 to 13 experiments.
Effect of wortmannin on binding of SP-A to alveolar macrophages. The assay was done according to Pison et al. (31). Briefly, macrophages in PBS were seeded onto 24-well plates at a density of 2.5 × 105 cells/well and then exposed for 1 h at 37°C to 0 or 100 nM wortmannin. Afterward, the cells were cooled, washed three times with cold DMEM containing 0.1% bovine serum albumin (binding buffer), and incubated for 6 h at 4°C with trace amounts of labeled SP-A and increasing amounts (0.5-25 µg/ml) of unlabeled SP-A. After the incubation, the cells were washed, lysed, and used to measure protein (32) and for counting.
Specific binding was defined as total binding minus binding measured in the presence of a 100 times excess of unlabeled SP-A (nonspecific binding). Treatment with wortmannin did not change nonspecific binding (data not shown). Each assay was done in triplicate, and the experiment was repeated three times. The results are presented as nanograms of SP-A bound per microgram of cell protein.Effect of wortmannin on the internalization of bound SP-A. In this assay, macrophages in RBA were seeded onto six-well plates (2 × 106 cells/well), allowed to adhere for 2 h at 37°C, and then incubated for 1 h with 0 or 100 nM wortmannin in RBA. Afterward, the plates were placed on ice, and the cells were incubated for 30 min with cold RBA containing 1 µg/ml of 125I-SP-A. The ligand was then removed, and the cells were rinsed three times with cold RBA. At this point, reference cells were incubated for 3 min at 37°C with a mixture of 0.035% trypsin and 0.013% EDTA in 15 mM phosphate-citrate saline, pH 5, and the medium was harvested and counted. The cells on which internalization of SP-A was measured were first incubated for 30 s with RBA at 37°C to allow the partial internalization of bound ligand and then exposed for 3 min to trypsin-EDTA at 37°C to detach SP-A still bound to the plasma membrane. These cells were then harvested by scraping, pelleted, and used to count radioactivity and to measure protein. The radioactivity associated with the cell pellet is expressed as a percentage of the radioactivity liberated from reference cells. Experiments were done in triplicate and repeated four times. In preliminary experiments, we found that control macrophages internalize 55 ± 17% (SE) of bound SP-A in 30 s (n = 4 experiments).
Effect of wortmannin on steps after internalization of SP-A. Macrophages were seeded onto six-wells plates (2 × 106 cells/well) and then incubated with 125I-SP-A (1 µg/ml RBA) for 15 min at 37°C. Afterward, labeled SP-A was removed, and the cells were incubated at 37°C for 30 min with 0 or 100 nM wortmannin in RBA. After the incubation, TCA-soluble radioactivity was measured in the medium and cell homogenate. Data presented are from three experiments.
Effect of wortmannin on the association of SP-A with organelles
isolated from the postnuclear supernatant.
Alveolar macrophages in RBA were seeded onto six-well plates (2 × 106 cells/well) and then incubated for 1 h with 0 or
100 nM wortmannin in RBA. 125I-SP-A (1 µg/ml RBA) was
then added, and the incubation was continued for 15 min at 37°C.
Afterward, the medium was removed, and the cells were first rinsed
three times with RBA and then incubated in RBA for a further 30 min at
37°C (chase). The cells were then homogenized in 0.25 M sucrose and 1 mM EDTA, pH 7.4, and the homogenate was centrifuged at 750 g
for 7 min to obtain a postnuclear supernatant. Aliquots of the
postnuclear supernatant (1.5-2 ml obtained from 1 × 106 to 2 × 106 cells) were deposited over
a discontinuous gradient made by a 1.2-ml cushion of 2.5 M sucrose and
1 mM EDTA, pH 7.4, and an intermediate layer of 18% Percoll in
homogenization buffer (8.5 ml) and were centrifuged for 30 min at
20,000 rpm in a Ti 50 rotor (Beckman, Palo Alto, CA) (23).
Then, the gradient was fractionated starting from the bottom, and the
fractions were used to count total and TCA-soluble radioactivity and to
measure -hexosaminidase (a lysosomal marker) activity
(23). When isolated according to this protocol, lysosomes
migrate toward the bottom of the tube (23), endosomes
remain at the top (14, 23), and SP-A is recovered almost quantitatively in the top 10 fractions. Results shown
are representative of three experiments.
Phagocytosis of opsonized sheep erythrocytes. Before use, sheep erythrocytes (109/ml PBS) were washed with PBS, incubated for 20 min at 37°C with rabbit IgG anti-sheep erythrocytes (80 µg IgG/ml), washed again, and suspended in DMEM so that 200 µl contained 107 opsonized erythrocytes.
Rabbit alveolar macrophages in DMEM were plated onto 24-well plates (2.5 × 105 cells/well) the day before the experiment; switched to DMEM supplemented with 10% heat-inactivated fetal calf serum, 50 ng/ml of LPS, 50 U/ml of penicillin and 50 µg/ml of streptomycin; and incubated overnight. Incubation with LPS was necessary to increase the baseline phagocytosis of red blood cells. On the day of the experiment, the macrophages were incubated for 1 h with DMEM containing 0 or 100 nM wortmannin. Afterward, 200 µl of opsonized erythrocytes were added to each well, and the incubation was continued for 2 h at 37°C to allow phagocytosis. Then, the plates were gently decanted, dipped in distilled water for 30 s to rupture the red blood cells bound to the plasma membrane, and rinsed three times with 1 liter of PBS. The macrophages were fixed (3) and analyzed with an inverted phase-contrast microscope, and the number of erythrocytes per 100 macrophages was counted as the phagocytic index. Each experiment was done in triplicate. Data are from four experiments. Under the present conditions, the phagocytic index of control macrophages was 366 ± 53 (SE; n = 4 experiments).Fluid-phase pinocytosis. Briefly, alveolar macrophages in RBA were allowed to adhere to 24-well plates for 2 h at 37°C (3 × 105 to 6 × 105 cells/well) and then were exposed for 1 h to 0 or 100 nM wortmannin in RBA. The medium was then changed to RBA containing 0.5 mg/ml of fluorescein dextran with average molecular weight of 150,000 (FD150), and the macrophages were incubated at 37°C for 2 more hours. After the incubation, the plates were drained, rinsed three times for 5 min in 1 liter of cold PBS, and drained again. The well contents were then lysed for 1 h at 37°C with 1 ml of 0.1% Triton X-100 and 50 mM Tris, pH 8.5, and fluorescence of the lysate was measured with a spectrofluorimeter (PerkinElmer RS 3B) with excitation at 495 nm and emission at 514 nm (3). Protein concentration in the lysates was also measured, and the results are expressed as fluorescence per microgram of cell protein. Each experiment was done in triplicate. Data are from three experiments.
Preparation of liposomes. We prepared three different sets of liposomes. Liposomes A contained (by weight) 50% DPPC, 35% egg PC, 10% PG, and 5% Chol. Liposomes B contained 48% DPPC, 32% egg PC, 5% PG, 10% PS, and 5% Chol. Liposomes C, prepared by extracting a commercial porcine surfactant (Curosurf, Chiesi Farmaceutici, Parma, Italy) according to Bligh and Dyer (9), contained 98% lipids and 1% each of SP-B and SP-C.
To prepare the liposomes, the lipids were evaporated in a round-bottom flask, mixed with the appropriate amount of [3H]DPPC, and dried again. The dried lipids were hydrated in PBS at room temperature for 1 h and then sonicated five times for 20 s each at 35 W. The final suspension was stored at 4°C and used within 3 wk.Degradation of DPPC by alveolar macrophages. Macrophages (2.5-5 × 106/ml) in RBA were incubated at 37°C for 1 h in the absence and presence of various drugs and for 1 more hour with liposomes containing [3H]DPPC (final concentration 50 µg of total lipids, 0.05 µCi/ml). The reaction mixture was then extracted according to Bligh and Dyer (9) in the presence of 0.2 mg of rabbit surfactant (carrier), and one aliquot of the water-methanol phase was counted in the presence of Hyonic Fluor (Packard). To calculate the degradation of DPPC by macrophages, the spontaneous degradation was subtracted. The degradation of DPPC by the conditioned medium was also studied. The experiments were replicated 3-14 times. The results are presented as picomoles of DPPC degraded per 106 cells per hour.
Analysis of data. Results are presented as means ± SE. Differences between groups were analyzed by ANOVA. The effect of the drugs on various macrophage activities is expressed as a percentage of the activity measured under control conditions, and the difference was analyzed by paired t-test. The accepted level of significance was 5%.
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RESULTS |
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Degradation of SP-A by alveolar macrophages.
In agreement with previous findings (6, 40),
we found that alveolar macrophages take up SP-A in a time-dependent
fashion, degrade it, and deliver the degradation products to the
extracellular milieu. In fact, during incubation of
125I-SP-A with the macrophages, the cell-associated
radioactivity increased from 0 to 20 min (P < 0.05 by
ANOVA) and then remained unchanged (Fig.
1, control cells). Furthermore, the
macrophages generated degradation products of SP-A that were mostly
recovered from the medium (Fig. 2,
control cells). We also found that, after 100 min of incubation, SP-A
bound to the plasma membrane represented 5 ± 1% of the
cell-associated radioactivity.
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Identification of cell structures involved in the
degradation of SP-A.
As shown in Fig. 3, the
degradation of SP-A by alveolar macrophages was not affected by LPS or
IFN-, known stimulants of phagocytosis and fluid-phase pinocytosis
(3, 12), or by filipin, a drug that
sequesters membrane Chol and interferes with invagination and coat
assembly of caveolae (2). In agreement with the last observation, methyl-
-cyclodextrin, a depletor of cell cholesterol and inhibitor of caveolae-mediated uptake (2), also had no effect on the degradation of SP-A (data not shown). On the other hand,
chlorpromazine, which inhibits endocytosis through clathrin-coated vesicles (39), greatly decreased the degradation of SP-A
(Fig. 3).
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Influence of PI3K on the degradation of SP-A. Inhibition of PI3K had striking effects on the way macrophages interact with SP-A. In fact, wortmannin inhibited both the association of SP-A with alveolar macrophages (Fig. 1) and the generation of TCA-soluble by-products (Fig. 2). As shown in Fig. 1 the effect of wortmannin on the association of SP-A with macrophages was evident during the first 60 min of incubation, whereas at 100 min, there was no difference from control cells. Interestingly, 50 µM LY-294002 inhibited the degradation of SP-A to an extent comparable to that of 100 nM wortmannin (Fig. 3), suggesting that the observed effects are most likely due to the inhibition of class IA or class IB PI3K. In fact, the inhibition of class II PI3K requires higher concentrations of wortmannin (38), and class III PI3K is not inhibited by LY-294002 (16). Our results indicate also that inhibition of PI3K did not induce an intracellular accumulation of SP-A degradation products (Fig. 2).
Because PI3K could influence the degradation of SP-A at several steps, we analyzed the effect of wortmannin on the binding of SP-A to the plasma membrane, internalization of bound SP-A, and processing of internalized SP-A. Binding of SP-A to rabbit alveolar macrophages was very similar to binding to rat macrophages as measured by Pison et al. (31) and was not significantly modified by wortmannin (Fig. 4). On the other hand, wortmannin slowed the internalization of SP-A bound to the plasma membrane (Table 1). The effect on the internalization of SP-A was different from that on the uptake of opsonized red blood cells or FD150. In fact, in the presence of wortmannin, the uptake of SP-A decreased to 63 ± 8% of the control value, whereas the uptake of red blood cells and FD150 decreased to 5 ± 2 and 6 ± 5%, respectively, of the control value (Table 1). The internalization of SP-A appears to be quick because control macrophages internalized 55 ± 17% of bound SP-A in 30 s (n = 4 experiments). This could explain why, while studying the association of SP-A with macrophages (Fig. 1), we found that wortmannin inhibited the association even at time 0 when the cells were exposed to SP-A for 45-60 s before being washed and counted.
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Degradation of DPPC by alveolar macrophages.
Under the present conditions, macrophages degraded 239 ± 36 pmol
DPPC · 106
cells1 · h
1 (n = 14 experiments), in agreement with the findings of Wright and Youmans
(40). The process is temperature dependent because the
degradation dropped to 9 ± 4% of the control value
(n = 4 experiments) when the incubation was done at
4°C. We also found that the conditioned medium contributed to 8 ± 4% of the observed degradation when, as in this experiment,
liposomes contained DPPC labeled in palmitate position 2 (n = 4 experiments). However, if the DPPC label was
located in the choline moiety, the contribution of the conditioned
medium to the overall degradation became 52 ± 13% of the total
value (n = 10 experiments).
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DISCUSSION |
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Degradation of SP-A and DPPC by alveolar macrophages. Resident alveolar macrophages are exposed to different surfactant forms: precursors of the interfacial film, the film itself, products generated by perturbations of the interface, and fractions generated by the interaction of surfactant with dusts, infectious agents, or alveolar waste products (4). The mechanism that directs only a few of these potential substrates toward the macrophages is still unclear. This experiment analyzed specifically how alveolar macrophages degrade SP-A presented in soluble form and DPPC presented as small liposomes.
Our results indicate that SP-A binds to the plasma membrane, is rapidly internalized, and is then moved toward degradative compartments. We also found that an intact cytoskeleton and an acidic milieu are necessary for the degradation of SP-A and that PI3K regulates the speed of degradation of SP-A by acting at several steps. Finally, it appears that the turnover time of SP-A taken up by alveolar macrophages, <100 min in basal conditions, can be modulated. On the other hand, our data indicate that DPPC is degraded through different mechanisms that seem to lack the flexibility displayed by those utilized during the degradation of SP-A. In this experiment, SP-A and DPPC were presented to alveolar macrophages at concentrations of 1 and 25 µg/ml, respectively, to reproduce the ratio found in BAL fluids. Considering that 106 macrophages degraded 2.4 ± 0.6% of SP-A and 0.7 ± 0.1% of DPPC in 1 h, it appears that, on a molar basis, the propensity of macrophages to degrade SP-A is several orders of magnitude greater than that to degrade DPPC. This observation supports the view that SP-A might behave as a housekeeping molecule in the alveoli and macrophages as the sites where materials bound to SP-A are continually checked and degraded.Cell structures and activities involved in the degradation of SP-A.
The techniques used here were not specifically designed to analyze the
mechanism of internalization of SP-A; however, the available evidence
suggests that this process might be clathrin mediated, whereas
phagocytosis, pinocytosis, or uptake through caveolae is less likely.
In fact, stimulators of phagocytosis, such as LPS or IFN-, or
inhibitors of the formation of caveolae, such as filipin or
methyl-
-cyclodextrin, had no effect on the degradation of SP-A. On
the other hand, chlorpromazine, an inhibitor of clathrin-mediated
uptake, had a powerful inhibitory activity. That phagocytosis and
pinocytosis have minor importance for the internalization of SP-A is
also suggested by the fact that wortmannin slowed the internalization
of SP-A bound to the plasma membrane but blocked the uptake of
opsonized red blood cells and FD150. In agreement with our findings and
interpretation, Araki et al. (3), using murine bone
marrow-derived macrophages, found that treatment with 100 nM wortmannin
blocked the uptake of opsonized red blood cells and FD150 but only
slightly inhibited the clathrin-mediated uptake of acetylated
low-density lipoproteins.
Mechanism of degradation of DPPC. DPPC, presented as small liposomes, is taken up and degraded by alveolar macrophages through mechanisms different from those utilized for the degradation of SP-A. In fact, substances that affect phagocytosis and pinocytosis, such as LPS or wortmannin, or inhibit caveolae-mediated uptake, such as filipin, had no effect on the degradation of DPPC. Similarly, perturbants of the cytoskeleton or dissipators of the lysosomal acidic milieu had no effect. Taken together, these findings suggest that DPPC might enter the macrophage by fusion with the plasma membrane and might be subsequently hydrolyzed through a process that does not require an active role for the cytoskeleton or an acidic lysosomal milieu. Chlorpromazine increased the degradation of DPPC by alveolar macrophages. However, from the present data, it is unclear whether the effect was due to increased uptake or accelerated transport toward the sites of degradation because chlorpromazine can affect the interaction of cells with lipophilic ligands both by influencing the physical properties of plasma membrane (19) and by favoring the delivery to specific intracellular sites. For example, it has been shown that chlorpromazine increases the delivery of Rhodobacter sphaeroides endotoxin from the cell periphery to perinuclear vesicular organelles (19).
The mechanism by which cytochalasin D increased the degradation of DPPC remains unclear. Increasing in two ways (by the presence of SP-A in the medium or PS in the liposomes) the association of lipids with the macrophages did not increase the degradation rate of DPPC. This lack of flexibility suggests that the macrophages could play a minor role in the regulation of the alveolar levels of surfactant remnants made by small vesicles (the main catabolic form of surfactant lipids) (4). On the other hand, the present results do not negate the possibility that macrophages could play an important role in the removal of large by-products of surfactant, such as those generated by the collapse of the interface or the interaction of surfactant with exogenous materials like dusts, infectious agents, or alveolar waste products. To summarize, SP-A and DPPC are degraded by alveolar macrophages through different pathways. Disturbances along each of these pathways could lead to distinct lung disorders. ![]() |
ACKNOWLEDGEMENTS |
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We thank Prof. Gaetano Crepaldi for continuous support, Prof. Alessandro Bruni for critical reading of the manuscript, and Prof. Fulvio Ursini for useful advice. We also thank Dr. Martina Zaninotto, Dr. Carlo Spirlì, and Raffaella Marin for help with the biochemical analysis.
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FOOTNOTES |
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Address for reprint requests and other correspondence: A. Baritussio, Clinica Medica I, Policlinico, Via Giustiniani 2, 35128 Padua, Italy.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Received 17 May 1999; accepted in final form 21 February 2000.
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