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Department of Cell Biology, Duke University Medical Center, Durham, NC 27710
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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125%. However, other
proteins that are structurally homologous to SP-A, mannose-binding
lectin and complement protein 1q, did not. SP-A enhances phagocytosis
via an opsonization-dependent mechanism and binds apoptotic PMNs
4-fold more than viable PMNs. Also, binding of SP-A to apoptotic
PMNs does not appear to involve SP-A’s lectin domain. These data
suggest that the pulmonary collectins SP-A and SP-D facilitate the
resolution of inflammation by accelerating apoptotic PMN
clearance. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Both soluble factors and phagocyte membrane receptors have been
identified that recognize apoptotic cells and mediate their clearance
(reviewed in Ref. 9). Soluble mediators that have been
identified include 
2-glycoprotein
(10) and complement proteins, which becomes deposited on
the apoptotic cell surface (11). Also, thrombospondin acts
in concert with the phagocyte receptors
v3 and CD36 to bridge
the apoptotic cell with the phagocyte (12). In addition to
v3 and CD36, other
phagocyte receptors that have been identified are the LPS receptor
CD14, class A and class B scavenger receptors, and an ATP-binding
cassette transporter, ABC-1. Inhibition studies suggest that an
unidentified phosphatidylserine receptor and lectin(s) are also
involved. As discussed in the review by Savill (13),
multiple interactions exist between the apoptotic cell and its
phagocyte.
Immunoregulatory proteins have been identified in the lungs. Among these proteins are surfactant proteins A and D (SP-A and SP-D), which are in the collectin protein family of innate immune molecules. Collectins are oligomeric proteins characterized by N-terminal collagen-like domains and C-terminal lectin or carbohydrate binding domains (14, 15). All family members are pattern recognition molecules that bind nonself moieties, thereby targeting the foreign material for rapid clearance by immune cells. SP-A in particular can bind to a variety of substrates, including carbohydrates (16), surfactant lipids and lipid vesicles (17, 18), and proteins (i.e., myosin (19)). Binding of SP-A to both bacteria and lipid vesicles enhances their phagocytosis by macrophages (20, 21, 22, 23, 24). The macrophages with which SP-A and SP-D interact are the resident immune cells in the lungs, the alveolar macrophages (AM).
Several in vivo models of pulmonary injury demonstrate the importance of AM phagocytosis of apoptotic PMNs in the resolution of inflammation (25, 26), and it has been suggested that chronic pulmonary inflammation may be a result of inefficient clearance of apoptotic PMNs (27). We hypothesize that when PMNs undergo apoptosis, the changes that occur at the PMN plasma membrane allow SP-A and SP-D to recognize them as nonself. The proteins would then bind the apoptotic PMNs and target them for rapid phagocytosis by AMs. In this way, SP-A and SP-D would protect the pulmonary tissue by quickly resolving inflammation.
To test this hypothesis, we used a well-established assay for detecting macrophage phagocytosis of apoptotic PMNs (2) and showed that SP-A and, to a lesser extent, SP-D accelerate AM phagocytosis of apoptotic PMNs. We then designed a more sensitive flow cytometric assay and showed that SP-A’s effect was time, dose, and opsonization dependent. The effect was not, however, affected by heat treatment of SP-A, but it was partially dependent on SP-A’s N-linked carbohydrates. Furthermore, SP-A binds apoptotic PMNs significantly more than viable PMNs, and this binding does not appear to involve SP-A’s carbohydrate recognition domain. Neither the structurally homologous serum collectin mannose-binding lectin (MBL) nor complement protein 1q (C1q) stimulated PMN uptake. Also, SP-A’s effect was specific to AMs, as peritoneal macrophages did not show enhanced phagocytosis of apoptotic PMNs in the presence of SP-A.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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All chemicals used were from Sigma (St. Louis, MO) unless otherwise indicated. PBS, RPMI 1640, DMEM, and IMDM were obtained from Life Technologies (Grand Island, NY). Jurkat cells were obtained from American Type Culture Collection (Manassas, VA). The fluorescent labels FITC, Alexa Fluor 488, chloromethylbenzamido derivative of 1,1'-dioctadecyl-3,3,3',3'- tetramethylindocarbocyanine perchlorate (CM-DiI), and CellTracker Blue (CTB) were obtained from Molecular Probes (Eugene OR). Apoptosis detection reagents (PE-labeled annexin and 7-aminoactinomycin D) and anti-CD95 Ab were purchased from PharMingen (San Diego, CA).
Animals
Male Sprague Dawley rats were obtained from Charles River
(Raleigh, NC) or Taconic Farms (Germantown, NY). Rats weighed
200–400 g.
Protein purification
SP-A was purified from the bronchoalveolar lavage fluid of patients with alveolar proteinosis as previously described (28). Briefly, SP-A was extracted from lavage fluid with butanol and sequential solubilization in octylglucoside and 5 mM Tris, pH 7.4. The SP-A was treated with polymyxin B agarose beads to reduce endotoxin contamination. All SP-A preparations had <0.1 pg endotoxin/µg SP-A by the Limulus amebocyte lysate assay QCL-1000 (BioWhittaker, Walkersville, MD). SP-A was stored in 5 mM Tris, pH 7.4, at -20°C.
SP-D was purified from rats treated with silica to induce surfactant accumulation as previously described (29). Briefly, 4 wk after rats received an intratracheal administration of silica, their lungs were lavaged, and SP-D was purified from the lavage by ultracentrifugation and affinity and gel filtration chromatography. Purified SP-D was dialyzed into PBS and stored at 4°C.
MBL was purified from rat serum (Pel-Freeze, Rogers, AR) by affinity and gel filtration chromatography as previously described (30).
Human serum C1q was purchased from Advanced Research Technologies (San Diego, CA).
Modification of SP-A
The N-linked oligosaccharides of SP-A were
enzymatically removed by treatment with N-glycosidase F
(Calbiochem, La Jolla, CA) using a slight modification of a published
protocol (31). SP-A deglycosylation was conducted at
37°C for 16 h in 20 mM sodium phosphate (pH 7.4), 6 mM
n-octyl--D-glucopyranoside, and 10
mM EDTA with 1 µg enzyme/25 µg SP-A. The reaction mixture was
buffer exchanged into 10 mM sodium phosphate buffer, pH 7.4, using a
Centricon-100 concentrator (Amicon, Beverly, MA). Mock-treated SP-A was
prepared exactly as described above, except N-glycosidase F
was omitted. Gel electrophoresis was used to confirm that the SP-A was
completely deglycosylated. Final protein concentrations were determined
with the bicinchoninic protein assay kit (Pierce, Rockford,
IL).
SP-A was labeled with the amine-reactive probe FITC or Alexa Fluor 488 according to the manufacturer’s instructions. Briefly, 500 µg of SP-A was dialyzed into 10 mM sodium borate, pH 9.5, labeled with 50 µg of FITC for 1 h or dialyzed into 5 mM HEPES, pH 7.0, then labeled with Alexa Fluor 488 for 1 h. Excess FITC label was removed by dialysis against 5 mM Tris, pH 7.4, and excess Alexa label was removed by dialysis against 5 mM HEPES, pH 7.0.
Cell isolation
AMs were obtained by lung lavage of rats killed by sodium pentobarbital injection. Lungs were lavaged five to seven times with 10 ml of PBS and 0.2 mM EGTA. Cells were collected by centrifugation for 10 min at 330 x g and resuspended in the appropriate buffer. Macrophages were generally >98% pure as determined by hematoxylin differential stain.
Peritoneal macrophages were obtained by peritoneal lavage of rats 4–5 days after i.p. thioglycolate treatment (32). The peritoneum was lavaged five times with 10 ml PBS and 0.2 mM EGTA. Cells were collected by centrifugation for 10 min at 330 x g and resuspended in the appropriate buffer.
Peripheral PMNs were isolated as previously described with minor modifications (33, 34). Briefly, rats were anesthetized (initial experiments were performed with halothane/metofane, and later experiments yielding similar results were performed with i.m. injection of 0.1 mg of acepromazine, 7 mg of ketamine, and 29 mg of xylazine per 250 g rat), and the jugular was cannulated with a 23-gauge butterfly needle attached to a three-way stop cock. An exchange of 3–4 ml of 6% hetastarch solution with 100 U/ml heparin for 3–4 ml of blood was performed until 40 ml was exchanged per rat. The first 10 ml of blood was collected in a syringe containing 10 ml of the hetastarch solution. After the exchange, blood was collected until the rats died. Blood was allowed to settle 30–45 min at room temperature, and the RBC-depleted fraction was collected and centrifuged at 330 x g for 10 min. Cells were resuspended in 2 ml of PBS and underlaid with a five-step Percoll gradient. Step densities were 1.081, 1.085, 1.089, 1.093, and 1.097. Gradients were centrifuged at 500 x g for 30 min at room temperature, and the fractions at the interfaces of 1.085–1.089 and 1.089–1.093 density layers were pooled, washed, and resuspended in the appropriate buffer. PMN purities were generally >80% as determined by hematoxylin differential stain (EM Science, Gibbstown, NJ).
PMN apoptosis
Freshly isolated PMNs were induced to undergo apoptosis by 24-h culture or by UV radiation (10-min exposure, followed by 3- to 4-h culture) as previously described (35, 36). Apoptotic PMNs were then washed and suspended in the appropriate assay medium. The percent apoptosis was evaluated by PharMingen’s apoptosis detection kit using flow cytometry according to the manufacturer’s specifications. Freshly isolated PMNs were 27 ± 5% annexin positive, 24 h-cultured PMNs were 44 ± 12% annexin positive, and UV-irradiated PMNs were 80 ± 4% annexin positive.
Jurkat cell apoptosis
Apoptosis of Jurkat cells was induced by incubating them with anti-Fas mAb (37). Briefly, cells were cultured in RPMI 1640 and 10% heat-inactivated FCS and treated with 2 µg of anti-CD95 Ab/5 x 105 cells/ml for 25 h. Cells were then washed, and the percent apoptosis was evaluated by annexin staining. Jurkat cells were 38% annexin positive after Ab treatment.
Fluorescent labeling
Apoptotic PMNs were labeled with either CM-DiI for confocal microscopy or with CTB for flow cytometry. CM-DiI labeling was performed according to the manufacturer’s guidelines with slight modifications; PMNs were suspended at 2 x 106 cells/ml PBS, then treated with 0.75 µg of CM-DiI/106 cells for 10 min at 37°C followed by 15 min on ice. Cells were then washed and suspended at 2.5 x 107 cells/ml IMDM and 0.1% BSA.
PMNs were labeled with CTB by incubating cells in IMDM, 0.1% BSA, and 25 µM CTB for 40 min at 37°C in 5% CO2 followed by a second incubation for 30 min in fresh medium without CTB. Labeled PMNs were then suspended at 2.5 x 107 cells/ml IMDM in 0.1% BSA.
Macrophages were labeled according to manufacturer’s specifications in the PKH2 green fluorescent linker kit (Sigma). Briefly, macrophages were suspended at 2 x 107 cells/ml diluent A, then treated with an equal volume of 4 x 10-6 M PKH2 dye for 5 min at room temperature. To stop the labeling, an equal volume of 1% BSA in PBS was added to the mixture for 1 min. Cells were then washed with 1% BSA in PBS, followed by IMDM and 0.1% BSA, and suspended at 2 x 106/ml.
Myeloperoxidase-based phagocytosis assay
Uptake was evaluated based on methods previously described (1, 38, 39). Freshly isolated AMs were suspended at 1 x 106 cells/ml IMDM and 10% rat serum and adhered to 48-well tissue culture plates for 1 h at 37°C in 5% CO2. Wells were then washed twice with IMDM, and cultured PMNs were added to each well at 4 x 106 cells/ml IMDM with additional protein where indicated. Incubations were performed at 37°C in 5% CO2 for 30 min unless otherwise indicated. Wells were then washed three times with ice-cold PBS, fixed with 2% glutaraldehyde in PBS, and stained for myeloperoxidase (MPO) activity. To stain for MPO activity, the following was added to each well: 0.25 µl of HBSS, 0.1% BSA, 0.25 µl of 0.1 M phosphate buffer (pH 6.2), 10 µl of H2O2, and 10 µl of 1.25 mg/ml o-dianisidine. The reaction was conducted for 15 min at room temperature in the dark, then terminated with addition of 10 µl of 1% sodium azide. Control studies showed that AMs were 100% MPO negative, and PMNs were 100% MPO positive. Using light microscopy, cells were scored for the percentage of AMs that had PMNs associated with them. Sample wells were blinded, and a minimum of five fields of view at x100 objective and 500 macrophages were scored for each sample.
Flow cytometry phagocytosis assay
PKH2-labeled macrophages were incubated 1/3 with CTB-labeled apoptotic PMNs at a final concentration of 1.25 x 106 macrophages/ml IMDM and 0.1% BSA in BSA-precoated tubes in the presence of the indicated proteins. Cells were incubated at 37°C with gentle shaking for 60 min unless otherwise indicated. Cells were then washed with ice-cold PBS and 2 mM EDTA and fixed in 1% formaldehyde and 0.1% BSA in PBS for flow cytometry. Cells were analyzed for both labels, and PKH2-positive cells that were also positive for CTB were considered to have phagocytosed PMNs. Dual-labeled cells compared with total PKH2-positive cells were expressed as the percentage of macrophages that had phagocytosed PMNs.
SP-A binding assay
Cells were labeled using a slight modification of the protocol described in PharMingen’s Apoptosis Detection Kit. Briefly, cells were washed twice with ice-cold PBS, then suspended at 1 x 106 cells/ml binding buffer (10 mM HEPES/NaOH (pH 7.4), 0.14 mM NaCl, and 2.5 mM CaCl2). Aliquots (100 µl) were used for flow cytometry; cells were incubated with 25 µg/ml FITC-SP-A or Alexa-SP-A for 15 min on ice, then with 5 µl/tube PE-annexin for 15 min at room temperature. The binding reaction was stopped by adding 0.4 ml of ice-cold binding buffer/tube. Cells were analyzed by flow cytometry within 1 h or were fixed in 1% formaldehyde and stored at 4°C until analysis.
PMN studies measuring binding of different doses of SP-A and modified SP-A were performed as described, except that before labeling, annexin-positive cells were purified using the MACS apoptotic cell isolation kit (Miltenyi Biotec, Bergisch Gladbach, Germany) according to the manufacturer’s specifications.
Confocal microscopy
Phagocytosis was conducted as described above, except CM-DiI-labeled PMNs were used. After cells had been fixed, smears were made of 2 x 105 cells. Slides were air-dried, then treated with 10 µl of mounting medium (PBS, 50% glycerol, and 2% 1, 4-diazabicyclo[2.2.2]octane), covered with a coverslip, and sealed with nail polish. Cells were then examined under oil at x630 magnification. Images were captured using dual-channel excitation. The percent phagocytosis was determined by scoring AMs that had clearly internalized PMNs.
Statistical analysis
All data are expressed as the mean ± SEM, and statistical analysis was performed using one-tailed Student’s t test for unpaired samples.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We examined the ability of SP-A to enhance AM phagocytosis of
apoptotic PMNs using a well-established visual assay that detects
phagocytosed PMNs by staining for MPO activity (2). At
concentrations that stimulated phagocytosis of bacteria (21, 40), SP-A and, to a lesser extent, SP-D increased AM
phagocytosis of apoptotic PMNs (280 ± 62 and 158 ± 16% of
control value, respectively; Fig. 1
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6% of AMs contained detectable MPO),
and a time course was impossible to detect using the visual assay. We
suspected that AMs were rapidly degrading PMNs, and we therefore
designed a flow cytometric assay that allowed the detection of
phagocytosed PMNs even after they were broken down by AMs. PMNs and AMs
were labeled with different fluorescent dyes before phagocytosis. AMs
were then analyzed for PMN label. Dual-labeled AMs were considered
indicative of phagocytosis. These dyes have been designed for in vivo
studies, and cell-to-cell transfer is considered negligible during the
time course of these assays (41, 42).
Fig. 2 shows confocal microscopic images
of the individually labeled cells (A and B) and
images of AMs that had phagocytosed PMNs (C–F). The percent
AM phagocytosis of PMNs was determined by scoring confocal images to
confirm that the flow cytometric assay was an accurate measure of
phagocytosis and not just a measure of cell association. The flow
cytometric assay, like the MPO-based assay, showed that SP-A enhanced
AM phagocytosis of apoptotic PMNs, and this was confirmed by confocal
microscopy. However, baseline phagocytosis in these three assays was
significantly different, presumably due to the sensitivity of each
assay and the standard set for determining phagocytosis (MPO assay,
2.0 ± 0.3% phagocytosis; flow cytometry assay, 6 ± 1%
phagocytosis; confocal assay, 47 and 23% phagocytosis). Phagocytosis
in the presence of 25 µg/ml SP-A for the three assays was 3.7 ±
0.2% (MPO assay), 18 ± 3% (flow cytometry assay), and 61 and
53% (confocal assay).
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). Fig. 3A shows an example
of the flow analysis of PKH2-positive AMs for the CTB-PMN label. Fig. 3B is the dose-response curve of SP-A’s enhancement of
phagocytosis. Phagocytosis was increased in a dose-dependent manner,
with a plateau in the effect at 10–25 µg/ml.
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). SP-A’s effect peaked at 60 min, then
approached control levels.
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SP-A’s structural requirements for enhancement of phagocytosis
We examined the effect of modifying several aspects of SP-A’s
structure on its ability to enhance AM phagocytosis of apoptotic PMNs.
After boiling the protein for 10 min, there was no change in its
ability to enhance phagocytosis (Fig. 5).
However, when SP-A was deglycosylated, its ability to enhance
phagocytosis was significantly reduced (Fig. 5). Mock-treated SP-A
(SP-A exposed to the deglycosylation reaction but with no
N-glycosidase F) had an identical effect on phagocytosis as
untreated SP-A (data not shown).
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The ability of other collectins to stimulate phagocytosis of
apoptotic PMNs was examined. The lung collectin SP-D significantly
stimulated phagocytosis, although to a lesser extent than SP-A (Fig. 1
and Table I). The serum collectin MBL did
not significantly enhance phagocytosis at either 2 or 10 µg/ml (Table I). We also examined the ability of the structurally homologous C1q to
enhance phagocytosis, but no effect was seen (Table I).
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It has previously been shown that SP-A can stimulate AM
phagocytosis of bacteria by both coating the bacteria, thereby
targeting them for phagocytosis (opsonization-dependent phagocytosis)
(22, 23, 43), and directly stimulating AMs, thereby
causing them to be more phagocytically competent (activation
ligand-dependent phagocytosis) (21, 44). To determine the
mechanism by which SP-A enhances AM phagocytosis of apoptotic PMNs,
these two models were examined using an adherent visual assay. The
activation ligand-dependent model was tested by coating wells with SP-A
before AM adherence. Adhering AMs to SP-A-coated wells was not
sufficient to significantly increase AM phagocytosis of apoptotic PMNs
above the control value (Fig. 6).
However, when apoptotic PMNs were pretreated with SP-A for 15 min, and
unbound SP-A was removed by washing the PMNs, AMs phagocytosed
opsonized PMNs to the same extent as when SP-A was added directly to
the assay (Fig. 6).
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Macrophage phagocytosis of PMNs coincides with PMN apoptosis
(2, 39). Because SP-A seems to stimulate phagocytosis via
an opsonization-dependent mechanism (Fig. 6), and stimulation increases
as PMN apoptosis increases (data not shown), we hypothesized that SP-A
binding to PMNs increases as PMNs undergo apoptosis. To test this, we
compared SP-A binding to apoptotic vs viable PMNs. PMNs made apoptotic
by culturing were dual-labeled with FITC-SP-A and PE-annexin and
analyzed by flow cytometry. The intensity of SP-A binding to
annexin-positive (apoptotic) vs annexin-negative (viable) PMNs was
compared. Fig. 7 is a representative dot
plot of these experiments. SP-A bound to annexin-positive PMNs 4
± 1-fold more than to annexin-negative PMNs (average ± SEM;
n = 3). Annexin alone bound 84% of the PMNs (Fig. 7A), and SP-A alone bound 89% (Fig. 7B). Dual
labeling of the PMNs showed that annexin bound 69% and SP-A bound 86%
of the cells (Fig. 7C). This binding is dependent on SP-A
dose, but does not appear to be saturable at concentrations up to 25
µg/ml (Fig. 7D). An increase in SP-A binding to apoptotic
vs viable PMNs was also seen with the PMNs made apoptotic by UV
irradiation; SP-A bound annexin-positive PMNs 2-fold more than
annexin-negative PMNs. These data support the opsonization-dependent
model of SP-A-mediated AM phagocytosis of apoptotic PMNs. Binding of
modified SP-A to PMNs and AMs was compared to determine whether the
mechanism of binding to the two cells differed (Table II). Neither heat treatment nor
deglycosylation of SP-A significantly reduced SP-A binding to apoptotic
PMNs. However, both heat treatment and deglycosylation reduced SP-A
binding to AMs. The AM binding data are consistent with published
results by Oosting et al. (45).
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To determine whether SP-A has the capacity to bind other apoptotic cells, we induced apoptosis in Jurkat cells by cross-linking Fas on the surface of the cells (37) and compared SP-A binding to the apoptotic vs the viable cells. Apoptosis was detectable in 38% of the cells 25 h after induction, and SP-A bound apoptotic Jurkat cells 3-fold more than viable cells.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Collectins all bind to nonself molecular patterns. Before the current study the reported consequence of this binding was enhanced uptake of bacteria and viruses by phagocytes. The current study suggests that SP-A and SP-D can recognize apoptotic PMNs and enhance their phagocytosis by AMs. This effect was not shared by the serum collectin, MBL, or the structurally homologous complement component, C1q. Previous studies had demonstrated that C1q bound to apoptotic cells, but only in the presence of serum (46). As our experiments were performed without serum, it is not surprising that C1q was unable to enhance phagocytosis. It is possible that MBL may also require serum or other cofactors to enhance phagocytosis of apoptotic cells.
The ability of SP-A to enhance macrophage phagocytosis of apoptotic PMNs was not common to all phagocytes, as peritoneal macrophages did not show increased phagocytosis of apoptotic PMNs in the presence of SP-A. The peritoneal macrophages are, however, primed, because they have migrated into the peritoneum in response to an inflammatory stimulus. This priming may affect the macrophages’ ability to respond to SP-A, and it remains to be determined how the activation state of AMs may affect SP-A’s regulation of their phagocytosis of apoptotic PMNs.
SP-A enhanced AM phagocytosis in a time-dependent manner, with a peak in enhancement observed at 60 min. It was interesting that SP-A’s effect seemed to decrease after 60 min, although this is not statistically significant. This may be due to a decrease in the effective concentration of SP-A in the medium over the course of the assay, perhaps by AM uptake and degradation (47). Alternatively, an apparent reduction in phagocytosis may be due to a breakdown or dissipation of the PMN label after phagocytosis by the AMs, because fluorescence of the PMN label is dependent on its local concentration.
Multiple changes occur on the surface of apoptotic cells, revealing cryptic patterns that SP-A may then recognize as nonself. For example, changes in carbohydrates occur on the plasma membranes of apoptotic cells (5). Although the details of these changes remain to be determined, they appear to involve, at least partially, the loss of sialic acid residues on surface structures. Because mono- and disaccharides have been shown to inhibit phagocytosis of apoptotic cells (48, 49, 50), it has been speculated that a lectin receptor is involved in phagocytosis, and several lectin receptors on nonprofessional phagocytes have been identified (48, 50, 51).
It does not appear, however, that SP-A is interacting with the apoptotic PMNs via its lectin domain. Mannan and mannose, which bind to SP-A’s lectin domain, did not inhibit SP-A binding to the apoptotic PMNs. Also, the tertiary structure of this domain is what imparts the protein’s lectin ability; heat treatment of SP-A eliminates binding to immobilized D-mannose (16), attenuates enhancement of AM phagocytosis of bacteria (21), and reduces SP-A binding to AMs, which is consistent with our data (45). Interestingly, SP-A’s ability to mediate AM phagocytosis of apoptotic PMNs is not eliminated by heat treatment. This observation suggests that some domain other than the lectin domain is responsible for the interaction of the protein with apoptotic PMNs.
Although heat treatment of SP-A did not reduce phagocytosis or SP-A binding to apoptotic PMNs, it did significantly reduce SP-A binding to AMs. Studies attempting to characterize the SP-A receptor on AMs (reviewed by Tino and Wright (52)) suggest that SP-A has multiple interactions with AMs. Our data suggest that heat treatment of SP-A inhibits some of these interactions, but not those interactions required for mediation of AM phagocytosis of apoptotic PMNs.
An SP-A-mediated function that is not dependent on the higher structure of the protein is its ability to interact with lipid vesicles (17, 18). SP-A binds to a variety of lipids, including dipalmitoylphosphatidylcholine. Heat treatment of SP-A does not inhibit its ability to bind lipids, but it does decrease its ability to aggregate them. The only lipid change on the surface of apoptotic cells that has been identified is exposure of phosphatidylserine (3, 4). This phospholipid is actively partitioned to the inner leaflet of the plasma membrane in viable cells, but when cells become apoptotic, this asymmetry is lost, and phosphatidylserine is exposed on the surface of the cell. It seems unlikely that SP-A is binding to phosphatidylserine on apoptotic cells, because it has been previously shown that SP-A does not bind phosphatidylserine (18). It is possible that other unidentified changes in the phospholipid composition of the outer plasma membrane are responsible for SP-A binding or that SP-A may bind phosphatidylserine in the context of the plasma membrane.
Each monomer of SP-A is post-translationally modified by N-linked glycosylations. These glycosylations are not required for SP-A’s lectin activity or for binding to phospholipids (53), but they are involved in viral recognition (31, 54) and aggregation of phospholipid vesicles (17). Interestingly, SP-A’s N-linked carbohydrates are involved in AM phagocytosis of Mycobacterium tuberculosis (20). Also, we and others (45) have found that deglycosylation of SP-A significantly decreases SP-A binding to macrophages. The reduced ability of deglycosylated SP-A to enhance AM phagocytosis of apoptotic PMNs appears to be due to its reduced ability to bind AMs and not to a disrupted interaction with the PMNs.
SP-A not only bound apoptotic PMNs significantly more than viable PMNs, but it also bound apoptotic Jurkat cells significantly more than viable Jurkat cells. This suggests that SP-A is recognizing a motif on the apoptotic PMNs that is common to apoptotic cells regardless of cell type or mode by which apoptosis is induced. Identification of this binding site will not only contribute to a better understanding of how SP-A may regulate apoptotic cell clearance, but will contribute to our understanding of the changes that occur at the cell’s plasma membrane during apoptosis.
SP-A is not the first example of an innate immune molecule that plays a role in clearance of apoptotic cells. Both CD14, the primary LPS receptor, as well as complement have been found to mediate phagocytosis of apoptotic cells (11, 55). It seems possible that the clearance of bacteria and the clearance of apoptotic cells have evolved to encompass similar mechanisms. The goal of both systems is the same: to rid the body of unwanted cells or particles. Understanding how these proteins function will lead to a better understanding of how inflammation may be modulated to minimize potentially harmful proinflammatory responses while maximizing the host response against infection.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Jo Rae Wright, Box 3709, Department of Cell Biology, Duke University Medical Center, Durham, NC 27710.
3 Abbreviations used in this paper: PMN, polymorphonuclear neutrophil; SP-A, surfactant protein A; SP-D, surfactant protein D; AM, alveolar macrophage; C1q, complement protein 1q; MBL, mannose-binding lectin; CM-DiI, chloromethylbenzamido derivative of 1,1'-dioctadecyl-3,3,3',3'- tetramethylindocarbocyanine perchlorate; CTB, CellTracker Blue; MPO, myeloperoxidase.
Received for publication May 3, 2000. Accepted for publication November 27, 2000.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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RIII) expression. J. Immunol. 153:1254.[Abstract]
2-glycoprotein I in macrophage recognition. J. Biol. Chem. 272:31113. activity from LPS-stimulated macrophages. Am. J. Physiol. 15:L310.
, PGE2, and PAF. J. Clin. Invest. 101:890.[Medline]
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 Y. Hashimoto, T. Moki, T. Takizawa, A. Shiratsuchi, and Y. Nakanishi Evidence for Phagocytosis of Influenza Virus-Infected, Apoptotic Cells by Neutrophils and Macrophages in Mice J. Immunol., February 15, 2007; 178(4): 2448 - 2457. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. F. Vernon-Wilson, F. Aurade, and S. B. Brown CD31 promotes {beta}1 integrin-dependent engulfment of apoptotic Jurkat T lymphocytes opsonized for phagocytosis by fibronectin J. Leukoc. Biol., June 1, 2006; 79(6): 1260 - 1267. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M Bijl, E Reefman, G Horst, P C Limburg, and C G M Kallenberg Reduced uptake of apoptotic cells by macrophages in systemic lupus erythematosus: correlates with decreased serum levels of complement Ann Rheum Dis, January 1, 2006; 65(1): 57 - 63. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C.-H. Yang, J. Szeliga, J. Jordan, S. Faske, Z. Sever-Chroneos, B. Dorsett, R. E. Christian, R. E. Settlage, J. Shabanowitz, D. F. Hunt, et al. Identification of the Surfactant Protein A Receptor 210 as the Unconventional Myosin 18A J. Biol. Chem., October 14, 2005; 280(41): 34447 - 34457. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. E. Munoz, U. S. Gaipl, S. Franz, A. Sheriff, R. E. Voll, J. R. Kalden, and M. Herrmann SLE--a disease of clearance deficiency? Rheumatology, September 1, 2005; 44(9): 1101 - 1107. [Abstract] [Full Text] [PDF]
|
|
|
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