1 Institute for Environmental Medicine, University of Pennsylvania Medical Center, Philadelphia, Pennsylvania 19104; and 2 Department of Pediatrics and Cardiovascular Research Institute, University of California, San Francisco, San Francisco, California 94143
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ABSTRACT |
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The role of surfactant protein-A
(SP-A) in pulmonary uptake and metabolism of
[3H]dipalmitoylphosphatidylcholine
([3H]DPPC) was studied in SP-A gene-targeted mice
(SP-A /
). Unilamellar liposomes were instilled into the trachea of
anesthetized mice. Uptake was measured as dpm in lungs plus liver and
kidney for in vivo experiments and in lungs and perfusate for isolated
lung experiments. [3H]DPPC uptake increased with
CO2-induced hyperventilation in wild-type mice (SP-A +/+)
but was unchanged in SP-A
/
. Secretagogue treatment approximately
doubled the uptake of [3H]DPPC in isolated lungs from
SP-A +/+ but had no effect in SP-A
/
. Lungs degraded 23 ± 1.2% of internalized [3H]DPPC in SP-A +/+ and 36 ± 0.6% in SP-A
/
; degradation increased with 8-bromoadenosine
3',5'-cyclic monophosphate in SP-A +/+ but was unchanged in SP-A
/
.
Activity of lysosomal-type phospholipase A2
(PLA2) was significantly greater in lungs from SP-A
/
compared with SP-A +/+. Thus SP-A is necessary for lungs to respond to hyperventilation or secretagogues with increased DPPC uptake and also
modulates the PLA2-mediated degradation of internalized DPPC.
perfused lung; surfactant protein A knockout mice; lung surfactant; dipalmitoylphosphatidylcholine metabolism; lysosomal phospholipase A2
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INTRODUCTION |
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PULMONARY SURFACTANT is a highly conserved, heterogeneous mixture of lipids and proteins that is essential for normal lung function. Because either an excess or deficit of surfactant is deleterious, surfactant concentration is stringently regulated by pathways that involve synthesis, storage, secretion, and reuptake of surfactant lipid and protein components (9). A part of the internalized surfactant phospholipid is routed to lamellar bodies for resecretion, whereas the rest is degraded by lysosomes (9). To maintain a normal extracellular pool, the rate of surfactant clearance over the long term must equal the rate of secretion. Thus increased secretion due to hyperventilation or to administration of a secretagogue leads to increased clearance (11, 13, 26, 27).
Clearance of L--dipalmitoylphosphatidylcholine (DPPC) in
the intact lung is primarily by type II alveolar epithelial cells with
a contribution by alveolar macrophages (8, 9, 29, 34). The
use of inhibitors has suggested that endocytosis of DPPC by type II
cells occurs by both clathrin-dependent and clathrin-independent pathways (2, 25). However, the precise ligand for uptake via the clathrin-dependent pathway is not known. Because surfactant protein A (SP-A) binding to type II cells is clustered in coated pits
(30) and some in vitro studies have demonstrated a
SP-A-mediated increase in DPPC uptake (1, 33), it has been
proposed that the clathrin-dependent process occurs via
receptor-mediated uptake of a DPPC/SP-A complex (9,32).
However, comparison of wild-type (SP-A +/+) and SP-A gene-targeted
(SP-A
/
) mice showed similar rates of surfactant lipid clearance
(19) and similar alveolar phospholipid pool sizes
(20, 22). Because regulation of extracellular surfactant
pool size is crucial to survival, redundancy and excess capacity in the
clearance pathway might be expected. Thus we postulated that
SP-A-deficient mice might have normal basal uptake but might show
abnormalities in clearance when the rate of surfactant turnover is
increased during physiological stresses. To test this hypothesis, we
studied DPPC clearance in intact mice with CO2-stimulated
hyperventilation and in isolated mouse lungs perfused with
secretagogues. We utilized a newly developed SP-A gene-targeted mouse
as a model of SP-A deficiency.
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MATERIALS AND METHODS |
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Authentic lipids were obtained from Avanti (Birmingham, AL). Radiochemicals were from New England Nuclear (Boston, MA). 8-Bromoadenosine 3',5'-cyclic monophosphate (8-Br-cAMP), 12-O-tetradecanoylphorbol-13-acetate (TPA), bovine serum albumin (BSA), and fatty acid-free BSA were from Sigma (St. Louis, MO). 1-Hexadecyl-3-trifluoroethylgycero-sn-2-phosphomethanol (MJ33), a specific inhibitor of lysosomal-type phospholipase A2 activity in lungs (10), was a kind gift from Dr. Mahendra Jain, University of Delaware.
Generation of SP-A knockout mice.
The SP-A gene was targeted in F1 BL6/129Sv embryonic stem cells by
standard homologous recombination (17). Briefly, the replacement-type targeting vector replaced exons 2,
3, and 4, including the translation start site
for murine SP-A, and short segments of flanking intronic sequence.
Targeted clones were injected into day 2.5 postcoital
eight-cell to morula-stage CD1 zygotes and transferred to
pseudopregnant B6D2 females. Chimeric offspring were bred with albino
CD1 female mice, and their pups were screened by PCR and Southern
analysis for germline transmission of the mutant allele. Heterozygous
BL6/129Sv-CD1 mice were backcrossed onto a C57BL6 background. Mice used
for the initial part of this study were the 6th backcross generation
bred at University of California San Francisco. Subsequent experiments
used mice from the 10th backcross generation that were the progeny of a
breeding colony set up at the University of Pennsylvania. Analysis of
SP-A /
mice confirmed the absence of the normal SP-A gene (Southern blot) and SP-A mRNA (Northern blot) in the lung and of SP-A protein in
the lung lavage (Western blot). Wild-type C57BL6 mice were obtained
from Jackson Laboratories (Bar Harbor, ME). All procedures utilizing
mice were approved by the Institutional Animal Care and Use Committees.
Preparation of liposomes. Stock solutions of DPPC, egg phosphatidylcholine (PC), egg phosphatidyl glycerol, and cholesterol were mixed in a molar ratio of 10:5:2:3 with trace amounts of [choline-methyl-3H]DPPC ([3H]DPPC) to prepare liposomes (10). The phospholipid mixture was evaporated to dryness under N2, resuspended in phosphate-buffered saline (pH 7.4), and mixed vigorously. The suspension was subjected to three cycles of freezing under liquid nitrogen followed by immediate thawing at 50°C. The phospholipid suspension then was extruded under pressure through a 100-µm pore-size filter. Liposomes were stored at 4°C and used within 1 wk of preparation.
In vivo uptake of DPPC by mice and effect of hyperventilation. Mice were anesthetized with intraperitoneal xylazine (5.2 mg/kg) plus ketamine (40 mg/kg), and liposomes (10 nmol of DPPC in 20 µl of saline) were instilled intratracheally via a Hamilton syringe. The instilled dose was corrected for dpm remaining in the syringe plus dpm accumulated by the trachea and major bronchi measured at the end of the experiment. This correction represented <10% of instilled dpm. After recovery from anesthesia (within 20 min), mice were put into an acrylic chamber that was continuously flushed with 5% CO2 in air. Mice treated similarly, but allowed to breathe room air, were used as controls. After 3 h, mice were anesthetized with pentobarbital sodium (50 mg/kg body wt), and liver, kidney, and lungs were removed. Lungs were lavaged five times with 1-ml aliquots of saline, and lavage was collected. The trachea and major bronchi were separated from the lung parenchyma. The organs were then homogenized and counted for radioactivity. Uptake in tissues was calculated as the sum of dpm in lung, liver, and kidneys, and data were expressed as tissue dpm/lavage dpm.
In additional mice, ventilation parameters during air or CO2 breathing were measured using whole-body plethysmography (Buxco Electronics, Sharon, CT) (16). Briefly, each mouse was placed in the chamber, and the chamber pressure-time wave was measured continuously via a transducer connected to a computerized data-acquisition system. After baseline measurements for 30 min to ensure a steady state, frequency of breathing, tidal volume, and minute volume were recorded at 7.5-min intervals for 90 min during exposure either to air or 5% CO2.Isolated lung perfusion. Mice were anesthetized with intraperitoneal pentobarbital sodium (50 mg/kg body wt). [3H]DPPC-labeled liposomes (10 nmol of DPPC in 20 µl of saline) were instilled into the lungs through a cannula placed at the level of the tracheal carina. The mice were then continuously ventilated, the thorax was incised, and lungs were cleared of blood by perfusion through the pulmonary artery with buffer (Krebs-Ringer bicarbonate, pH 7.4, with 3% BSA). The total isolation procedure required ~5 min. Isolated lungs were perfused in a closed-circuit recirculating perfusion apparatus similar to that described previously for rat lungs (15). Perfusion buffer was the same as lung clearing buffer plus 10 mM glucose added. In some experiments, either 8-Br-cAMP (0.1 mM) or TPA (30 ng/ml) was added to the perfusate at the start of perfusion. Perfusate was gassed constantly with 5% CO2 in air. Lungs were ventilated at 60 cycles/min, 0.3 ml tidal volume, and 2 cmH2O end-expiratory pressure. There was no significant change in the ventilation and perfusion pressures or overt evidence of lung edema during the experiments.
DPPC uptake. At the end of the ~5-min period required for lung isolation (baseline) or the 2-h experimental perfusion, lungs were lavaged five times with 1-ml aliquots of ice-cold saline. Lung tissue was then homogenized in saline on ice with a Polytron homogenizer followed by a motorized mortar and pestle. Lung homogenate and perfusate aliquots were counted for dpm to calculate lung uptake of [3H]DPPC (dpm in the lung plus perfusate/total dpm instilled) (13). To calculate alveolar phospholipid pool size, we measured lipid phosphorous in the organic extract of lung lavage after cells were removed by centrifugation at 300 g for 10 min (10).
DPPC metabolism. Lipid and aqueous fractions from the lung homogenate were extracted by the Bligh and Dyer procedure (3). Phospholipids in the organic phase were fractionated by thin-layer chromatography on silica gel plates with chloroform-methanol-ammonia-water (65:35:2.5:2.5, vol/vol) as the solvent system (10), and bands of interest were scraped and counted for dpm. The disaturated phosphatidylcholine (DSPC) fraction was separated from total PC on a neutral alumina column after osmication of lipids (23). Unsaturated PC, representing PC in which fatty acyl components have one or more double bonds, was calculated as dpm in total PC minus dpm in DSPC. This fraction represents resynthesized PC, since all dpm were initially in DSPC (i.e., DPPC). Degradation of internalized DPPC was calculated from the sum of dpm in lung fractions (lysoPC, aqueous, and unsaturated PC) plus dpm in perfusate. This estimate does not include counts from DPPC metabolites that have been reincorporated into DSPC.
Lysosomal-type PLA2 activity of lung homogenates was measured in Ca2+-free, acidic (pH 4) buffer and also in pH 8.5 buffer in the presence of 10 mM Ca2+ (14). Substrate for both assays was liposomes with [3H]DPPC labeled in sn-2 palmitate as described previously (14).Statistical analysis. All results are expressed as means ± SE. Multiple group comparisons were done by one-way analysis of variance with Bonferroni's correction or with the Kruskal-Wallis method. Comparison of groups vs. a control was done using Dunn's method or Tukey's test using SigmaStat software version 2.0 (Jandel, San Rafael, CA). P < 0.05 was considered statistically significant.
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RESULTS |
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DPPC uptake in vivo.
We evaluated the alveolar uptake of DPPC and the effect of
hyperventilation in SP-A +/+ and SP-A /
mice. Recovery of total radioactivity at 3 h after instillation of 3H-labeled
liposomes was measured as the sum of dpm in lung lavage fluid, lung
tissue, liver, and kidney. Recovery for both SP-A+/+ and SP-A
/
mice varied from 68 to 98% of instilled dpm under control and
CO2 exposure conditions, indicating that a variable fraction of dpm ended up in other organs. There was no difference in
recoveries between control and CO2-stimulated conditions,
although recoveries in both conditions were slightly greater for SP-A
/
compared with SP-A +/+ mice (Table
1). In control SP-A +/+ mice, 55% of
instilled dpm were recovered in the lung lavage, 23% in the lung
tissue, and 13% in liver and kidney; the distribution of dpm in
control SP-A
/
mice was similar.
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DPPC uptake in isolated perfused lungs.
[3H]DPPC present in isolated lungs from SP-A +/+ mice at
the start of perfusion (5 min after instillation of
[3H]DPPC-labeled liposomes) was 4.4 ± 0.1% of
instilled dpm (n = 7) and was similar in SP-A /
mice (4.3 ± 0.09%, n = 3). This baseline uptake,
shown previously to represent primarily the rapid exchange of DPPC with
nonlavageable endogenous pools (11), was subtracted from
lung tissue values at subsequent time points. Baseline uptake is the
same for control and secretagogue-stimulated lungs, since it is
measured before the start of secretagogue infusion. Lung uptake of
[3H]DPPC during the subsequent 2 h of perfusion in
the absence of secretagogues was 5.1 ± 0.4% of instilled and was
comparable for SP-A +/+ and SP-A
/
mice (Fig.
2). However, the lungs from SP-A
/
mice responded differently from wild-type mice to secretagogues. With
SP-A +/+ mice, pulmonary uptake of [3H]DPPC was
significantly stimulated by 2.2-fold by treatment with secretagogues
(8-Br-cAMP, TPA), whereas secretagogues had no effect on uptake by
lungs from SP-A
/
mice (Fig. 2).
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Phospholipid pool size.
Phospholipid pool size was measured in the alveolar lavage fluid of
control mice and after 3 h of exposure to 5% CO2 and
also in the isolated lung after 2 h of perfusion. For intact mice, the measurement represents the endogenous pool in the absence of added
lipid, whereas measurements in the perfused lung were made at 2 h
after instillation of liposomes. The amount of phospholipid instilled
in liposomes represented ~5% of the endogenous pool. The pool size
was not different between SP-A +/+ and SP-A /
mice and did not
change significantly following treatment with CO2 exposure
or with secretagogues (Table 3).
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DPPC metabolism in isolated perfused lungs.
The distribution of dpm in total PC, unsaturated PC,
lysoPC, the aqueous fraction of the lung extract, and the
perfusate was determined to evaluate degradation of internalized DPPC
by the isolated perfused lung. The PC fraction (disaturated plus
unsaturated) accounted for ~90% of the total lung dpm in SP-A +/+
mice (Table 4). The unsaturated PC
fraction was ~13% of lung-associated dpm. This fraction represents
PC that is resynthesized from 3H-labeled DPPC degradation
products. Dpm in lysoPC and the aqueous lung extract, representing
[3H]DPPC degradation products, accounted for ~1 and
7%, respectively, of lung-associated dpm. Previous studies in rat
lungs showed that dpm in the aqueous fraction represent labeled
glycerophosphocholine, choline phosphate, CDP-choline, and free
choline, whereas dpm in the perfusate represent effluxed free choline
(12). Perfusate dpm accounted for only 1-2% of the
counts taken up by the lung in 2 h, and this value was not
altered significantly by the presence of secretagogues. Total
degradation of [3H]DPPC at 2 h, calculated as
the sum of dpm in aqueous metabolites (lung plus perfusate), lysoPC,
and unsaturated PC, was significantly greater (P < 0.05) in SP-A /
compared with SP-A +/+ mice (Table 4). 8-Br-cAMP
resulted in an increased rate of DPPC degradation by lungs from SP-A
+/+ mice as previously demonstrated for rat lungs (11),
but there was no effect of this secretagogue on DPPC degradation by
lungs from SP-A
/
mice (Table 4). Thus increased degradation with
8-Br-cAMP appears to reflect an increased DPPC uptake rather than a
direct effect on PLA2 activity consistent with previous
findings in rat and mouse lungs (11, 21).
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DISCUSSION |
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Surfactant phospholipid secretion responds to a wide variety of
mediators, including -adrenergic agonists such as 8-Br-cAMP and
protein kinase C agonists such as phorbol esters (4, 24, 31). Studies with these mediators in intact lungs or in alveolar type II cells in culture have shown that they also promote cellular uptake of phospholipid, thus giving rise to the concept that surfactant secretion and reuptake are linked (7, 9, 13). In support of this hypothesis, hyperventilation induced by exposure to
CO2 or by exercise stimulated both surfactant secretion as
well as reuptake (26, 27). Studies with isolated type II
cells in culture suggested that SP-A plays a role in surfactant
homeostasis, since the addition of SP-A to liposomes augmented cellular
uptake of DPPC (1, 33). Furthermore, SP-A was shown to
inhibit the lysosomal-type phospholipase A2, the enzyme
responsible for degradation of internalized DPPC (14).
However, SP-A gene-targeted mice had no significant derangement of DPPC
homeostasis under basal physiological conditions (18, 19).
These results do not exclude the possibility that the role of SP-A
might be critical when the rate of pulmonary surfactant turnover is
increased. The objective of the present study was to use genetically
transformed SP-A
/
mice to evaluate the role of SP-A in DPPC uptake
and metabolism under resting and stimulated conditions.
Hyperventilation with CO2 was chosen as a model for
stimulation of DPPC uptake in intact mice. Hyperventilation
significantly enhanced the uptake of intratracheally administered
[3H]DPPC by the lungs of SP-A +/+ mice as demonstrated by
decreased dpm remaining in the lung lavage and increased dpm in the
lung tissue plus distal organs (liver and kidney). Using a
PLA2 inhibitor, we demonstrated that liver and kidney dpm
arise from degradation of [3H]DPPC in the lung and are
presumably transported to the distal organs via the blood stream.
Although SP-A +/+ and SP-A /
mice exposed to CO2 had a
similar degree of hyperventilation, only the wild-type mice showed
increased DPPC uptake. These results suggest that SP-A plays a crucial
role in the increased DPPC uptake associated with the physiological
stress of hyperventilation.
We extended these investigations to in vitro conditions to more closely
regulate physiological parameters. The isolated perfused lung model has
been used extensively to investigate the synthesis and secretion of
alveolar phospholipids as well as to study clearance of phospholipid
from the alveoli (4, 10-13). Similar to previous results with rat lungs (11, 13), isolated perfused lungs
of SP-A +/+ mice exhibited increased uptake of DPPC in response to secretagogues (cAMP, TPA). On the other hand, SP-A /
mice showed no
change in DPPC uptake in the presence of secretagogues, confirming the
differences observed between wild-type and gene-targeted mice with in
vivo experiments.
Endocytosed DPPC may be either directly recycled or degraded with
subsequent reutilization of degradation products (9). Perfused lungs of SP-A /
mice showed a significantly higher level
of degradation of internalized DPPC under basal conditions compared
with wild-type controls. These results suggest that SP-A regulates
metabolism of internalized DPPC and are consistent with our previous
studies of isolated granular pneumocytes (14). These cells
showed a lower rate of degradation of PC in natural surfactant compared
with liposomes alone and a decrease in the degradation of liposomal
DPPC with added SP-A. This effect can be explained by in vitro
experiments showing that SP-A inhibits lysosomal-type PLA2
activity, the enzyme responsible for the initial step in degradation of
DPPC (14). Activity of lysosomal-type PLA2 was
significantly higher in lungs of SP-A
/
mice compared with the wild
type, compatible with a loss of PLA2 inhibition. As an
alternative mechanism, SP-A could direct intracellular trafficking of
the phospholipid-SP-A complex toward a nondegrading compartment in type
II cells (33), thereby promoting recycling of surfactant components. Thus the absence of endogenous SP-A could result in failure
of targeting and consequent increased rate of DPPC degradation.
The present results indicate the presence of two different mechanisms for DPPC uptake by lung cells. The first is an SP-A-independent mechanism that has sufficient activity to support basal levels of phospholipid recycling. The second is an SP-A-dependent mechanism that is required to maintain normal DPPC recycling rates under conditions of increased surfactant turnover. On the basis of previous results, these studies presumably reflect uptake by type II cells, since they are responsible for the major fraction of DPPC clearance (8, 9, 29, 34). However, alveolar macrophages or other cells may also participate. The SP-A-dependent pathway most likely involves uptake of an SP-A/DPPC complex. We have shown previously that SP-A-dependent uptake of DPPC in the type II cell occurs via a cell membrane SP-A receptor that can be recruited to the cell surface by secretagogues (6). Because SP-A binding has been localized to coated pits in type II cells (30), uptake of the DPPC/SP-A complex likely occurs via clathrin-mediated endocytosis, whereas SP-A-independent uptake may occur by another pathway, possibly involving cell membrane retrieval (2, 25).
The physiological implication of a failure to increase clearance with
hyperventilation or secretagogues would be a transient elevation of
alveolar phospholipid, assuming that secretory response to those
stimuli remains intact. A previous study of hyperventilation in SP-A
/
mice evaluated the DPPC pool size after 1 h of exercise by
running or swimming (18). In that study, alveolar DPPC
content after running was greater in SP-A
/
mice than for SP-A +/+
mice, although there was no difference following swimming exercise
(18). DPPC pool size also was similar in SP-A +/+ and SP-A
/
mice with hyperventilation or following treatment with 8-Br-cAMP
or TPA in the present experiments. Thus the regulation of alveolar DPPC
pool size appears to involve additional levels of control.
In conclusion, we have shown an increased lung uptake of [3H]DPPC in wild-type mice on exposure to a physical (hyperventilation) or chemical (secretagogue) stimulus. Increased [3H]DPPC uptake did not occur under those conditions in mice with genetic deficiency of SP-A. We propose that SP-A is necessary to maintain normal rates of DPPC clearance from the alveolar space under conditions of increased surfactant turnover.
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ACKNOWLEDGEMENTS |
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We thank Dr. Michael Beers for loan of the mouse plethysmograph and Michelle Sperry for excellent technical assistance.
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FOOTNOTES |
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This work was supported by National Heart, Lung, and Blood Institute Grants HL-19737, HL-24075, and HL-58047.
Address for reprint requests and other correspondence: A. B. Fisher, Inst. for Environmental Medicine, Univ. of Pennsylvania School of Medicine, 1 John Morgan Bldg., 3620 Hamilton Walk, Philadelphia, PA 19104-6068 (E-mail: abf{at}mail.med.upenn.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.
First published January 10, 2003;10.1152/ajplung.00200.2002
Received 24 June 2002; accepted in final form 17 December 2002.
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