Physiology Program, Harvard School of Public Health, Boston, Massachusetts 02115
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ABSTRACT |
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Although the tachykinins substance P (SP) and neurokinin A have been largely localized to neurons, eosinophils have also been shown to express these peptides. Our aim was to determine whether rat alveolar macrophages (AM) express preprotachykinin gene-I (PPT-I) mRNA that encodes these tachykinins and to examine expression during inflammation. PPT-I mRNA was detected by reverse transcription (RT)-polymerase chain reaction (PCR) in AM and brain (control) but not in peritoneal macrophages. Northern analysis showed that PPT-I mRNA was induced two- to fourfold by in vivo treatment of rats with intratracheal lipopolysaccharide (LPS) and in vitro after 4 h of exposure to LPS. This increase was inhibited by dexamethasone. In situ RT-PCR and immunocytochemistry further confirmed that AM express PPT-I mRNA and SP-like immunoreactivity, respectively, which was enhanced by LPS treatment. A 1.3-kb transcript consistent with PPT-I mRNA was detected by Northern analysis of bronchoalveolar lavage neutrophils. Therefore, rat AM express PPT-I mRNA that is upregulated in AM by LPS and is attenuated by dexamethasone. PPT-I mRNA was also detected in lung neutrophils.
substance P; neutrophil; lipopolysaccharide; bronchoalveolar lavage; lung
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INTRODUCTION |
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THE MAMMALIAN TACHYKININS substance P (SP), neurokinin
A (NKA), NKA-(310), neuropeptide K, and neuropeptide-
are produced as a result of differential splicing of a single transcript of the
preprotachykinin gene-I (PPT-I) or by posttranslational processing of
the peptides (17). These tachykinins participate in the regulation of
diverse biological processes in the central and peripheral nervous
systems, in endocrine glands, and in a variety of body systems,
including the respiratory tract. In particular, the undecapeptide SP
has been documented as both a neurotransmitter and an immunomodulator substance. SP can stimulate DNA and protein synthesis in T lymphocytes, evoke the release of inflammatory cytokines from blood monocytes, and
enhance the lysosomal enzyme release and phagocytic activity of human
neutrophils (2, 20). SP is also recognized as a chemical mediator of
neurogenic inflammation because it participates in vasodilatation,
increases vascular permeability, and affects cells involved in the
inflammatory and immune process (18).
The idea that the nervous system modulates immunological and inflammatory responses has been supported by the identification of neuropeptide receptors on leukocytes and the demonstration that these peptides can regulate leukocyte function. Mononuclear phagocytes, either as circulating blood monocytes or as tissue macrophages, influence host defense responses through their capacity to present antigens and to release a host of mediators (19, 20). SP can enhance phagocytosis by both neutrophils and macrophages (2) and can serve as a chemotactic agent to monocytes and macrophages (26). Blood monocytes have also been shown to release increased levels of interleukin (IL)-1 in the presence of SP (20), and, conversely, IL-1 can increase sympathetic neuron levels of SP (14). Thus there appears to be interaction between the immune system, the nervous system, and SP to modulate inflammation.
Neurons are the predominant, but not the exclusive, source of tachykinins. Eosinophils have been shown to synthesize immunoreactive SP (1) and also to express PPT-I mRNA (21). In situ hybridization performed on synovial tissue from rats with Freund's adjuvant-induced arthritis and humans with rheumatoid arthritis demonstrated PPT-I mRNA-positive cells that resemble macrophages (11) and a murine macrophage cell line that reportedly expresses SP (24). In addition, specific functional tachykinin receptors are present on human blood monocytes (20) and guinea pig alveolar macrophages (AM; see Ref. 4). In light of these scattered observations, we wanted to determine whether rat AM express PPT-I mRNA and to begin to examine the regulation of this message during lung inflammation. We demonstrated substantial expression of a single transcript size of PPT-I mRNA in rat AM. PPT-I mRNA was upregulated by lipopolysaccharide (LPS) exposure and was downregulated by dexamethasone pretreatment both in vivo and in vitro. SP-like immunoreactivity was evident in macrophages from both control and LPS-treated groups, but the fluorescence intensity was increased after LPS. Neutrophils collected by bronchoalveolar lavage (BAL) after intratracheal instillation of LPS also expressed PPT-I mRNA and appeared to be immunoreactive for SP.
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MATERIALS AND METHODS |
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Cell isolation and treatment.
Adult Sprague-Dawley rats were managed in accordance with the National
Institutes of Health standards defined by the United States Department
of Agriculture Animal Welfare Acts. Experimental protocols were
approved by the Harvard Medical Area Standing Committee on Animals.
Rats were euthanized with an overdose of pentobarbital sodium (65 mg
ip; Anthony Products, Arcadia, CA), and AM were collected by BAL with
10 5-ml washes of endotoxin-free Dulbecco's phosphate-buffered saline
(PBS) with 0.6 mM Na2-EDTA. Fluid
recovered from BAL was centrifuged (400 g) at 4°C. Viability (98%)
and total cell counts were determined by hemocytometer counts of
aliquots diluted in trypan blue solution. Cell type was determined from modified Wright-Giemsa-stained cytocentrifuge preparations (
95% AM).
Cells were cultured in RPMI 1640 with 10% fetal calf serum, 100 U/ml
penicillin, and 100 µg/ml streptomycin (Sigma Chemical, St. Louis,
MO) at 37°C in 5% CO2.
Macrophages were plated in 10 ml of medium at
106 cells/ml in 100-mm polystyrene
tissue culture plates (Fisher Scientific, Pittsburgh, PA) with or
without polyhydroxymethylacrylate coating to prevent cell adherence.
Cells were incubated with or without 10 µg/ml
Escherichia coli LPS (Sigma) for 4 h.
In some experiments, cells were incubated with
10
7 M dexamethasone (Sigma)
for 2 h before the addition of LPS (7). After incubation, cells were
prepared for total RNA extraction by lysis in 3 ml of modified
guanidine thiocyanate [GTC; 4 M guanidine thiocyanate, 25 mM
sodium citrate (pH 7.0), 0.5%
N-lauryl sarcosine, and 0.1 M
2-mercaptoethanol] and were stored at
70°C for RNA isolation. To determine if blood mononuclear cells contained PPT-I mRNA, blood was collected from some rats by intracardiac puncture and
was purified (>98% mononuclear cells) with the use of a
discontinuous two-step Nycodenz (Nycomed, Diagnostics Division, Oslo,
Norway) gradient (9). Cells were lysed in GTC and were stored at
70°C for RNA isolation. Rat brain (positive PPT-I mRNA
control) was also collected, homogenized in GTC solution with a
Brinkmann polytron, and stored at
70°C for RNA isolation.
RNA purification. Total RNA was extracted using a modified guanidinium method. The lysed cell mixture was thawed and layered on 5.7 M CsCl-0.1 M Na2-EDTA and centrifuged at 268,000 g for 4 h at 20°C in a Beckman SW55 Ti rotor. Pelleted RNA was resuspended in DEPC-treated 10 mM tris(hydroxymethyl)aminomethane (Tris) · HCl (pH 7.4) and 1 mM EDTA, ethanol precipitated, and resuspended in DEPC-treated water.
RT-PCR.
The PPT-I gene, from which both SP and NKA are produced, contains seven
exons, which, through alternative splicing, generate at least three
distinct PPT-I mRNAs (designated -,
-, and
-PPT-I; see Refs. 5
and 17). Primers used to amplify these transcripts corresponded to
sequences that span exons 2 and 3, which are common to all species of
PPT-I mRNA (Table 1). Total RNA (1.0 µg)
isolated from rat brain, peritoneal macrophages, or AM was reverse
transcribed with 17-base oligo(dT) as primer and 200 units Moloney
murine leukemia virus (MMLV) reverse transcriptase (Life Technologies, Gaithersburg, MD) at 37°C for 90 min. The cDNA was subjected to 30 cycles of amplification with 1.25 units AmpliTaq polymerase (Perkin-Elmer, Norwalk, CT) in 10 mM Tris · HCl (pH
8.3), 50 mM KCl, 1.5 mM MgCl2,
0.01% gelatin, 200 µM dNTP, and 1.0 mM each of the 5'- and
3'-primers (Table 1, set
1). Each cycle consisted of
denaturation at 94°C for 1 min, annealing at 55°C for 2 min, and extension at 72°C for 3 min.
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Probes.
An antisense PPT-I oligonucleotide for probing Southern blots was
5'-end labeled with T4 kinase (Pharmacia, Piscataway, NJ) and
6,000 Ci/mmol
[-32P]ATP (NEN,
Boston, MA). The PPT-I-specific antisense oligonucleotide utilized was
internal to the antisense oligonucleotide used during the PCR procedure
(Table 1).
Northern blotting and hybridization. Total RNA was denatured and resolved in 1.0% agarose-formaldehyde gels, transferred to Nytran filters, and UV cross-linked to the membrane. Prehybridization and hybridization were completed in 0.5 M NaPO4 (pH 7.2), 1 mM EDTA, 7% SDS, 150 µg of tRNA, and 10 ml of hybridization solution at 65°C. Blots were washed in 0.1× SSC-0.2% SDS one time for 15 min at 37°C and two times for 15 min at 40°C before autoradiography. In some experiments, quantitative densitometric scanning was performed.
In situ RT-PCR.
BAL cells collected 4 h after intratracheal instillation had been
dotted onto positively charged slides, fixed, and stored at
70°C as described above. The slides were treated with
proteinase K (6 µg/ml of PBS; Sigma) for 15-20 min, heated at
95°C for 2 min to inactivate the enzyme, rinsed in PBS and
DEPC-treated water, and air dried. The in situ RT and amplification
reaction solution contained each of the PPT-I-specific primers (0.2 µg; Table 1, set
2), 10 mM dNTP, 10 mM
Tris · Cl (pH 8.3), 50 mM KCl, 3.5 mM MgCl2, 80 units RNasin (Promega,
Madison, WI), 100 units MMLV reverse transcriptase (Life Technologies),
and 2.5 units AmpliTaq DNA polymerase (Perkin-Elmer). Slides were
incubated at 43°C for 1 h on the heating block of a programmable
thermal cycler (TwinBlock System; Ericomp, San Diego, CA). The
amplification reaction included 15 cycles of denaturation at 94°C
for 1 min, primer annealing at 60°C for 2 min, and extension at
72°C for 1 min. The amplification solution was removed, and the
slides were heated at 92°C for 1 min to immobilize the
intracellular signal. Slides were washed in 2× SSC for 5 min at
room temperature with gentle agitation. A PPT-I-specific 21-mer (Table
1, probe) was labeled by incorporation of a
digoxigenin-labeled nucleotide (DIG Oligonucleotide 3'-End Labeling Kit; Boehringer Mannheim, Indianapolis, IN). In addition to
the PPT-I-specific oligonucleotide and digoxigenin-labeled dideoxyuracil 5'-triphosphate, the probe mixture contained 5× reaction buffer [0.2 M potassium cacodylate, 25 mM
Tris · HCl (pH 6.6), 0.25 M bovine serum albumin, 5 mM CoCl2 solution, and 50 units
terminal transferase]. The mixture was incubated at 37°C for
15 min and then was placed on ice. Each 100 µl of hybridization solution contained 2 µl of digoxigenin-labeled oligonucleotide (0.2 µg/µl), 50 µl of deionized formamide, 10 µl of 20× SSC,
10 µl of 100× Denhardt's solution, 10 µl of 10 M herring
sperm DNA, 1 µl of 10% SDS, and 17 µl of deionized water. The
hybridization solution was added to the slides (50 µl/slide), heated
to 95°C for 5 min, and then maintained at 48°C for 4 h. After
hybridization, the slides were washed in 2× SSC with gentle
agitation for 15 min. The digoxigenin-labeled cDNA segments were
detected by an enzyme-linked immunoassay using an anti-digoxigenin
antibody conjugated with alkaline phosphatase (Boehringer Mannheim).
Slides and antibody solution were incubated overnight at 4°C. The
amplified gene product of interest was localized using an
enzyme-catalyzed color reaction with
5-bromo-4-chloro-3-indolyl phosphate (x-phosphate)
and nitro blue tetrazolium salt that produced an insoluble blue-black
precipitate after incubation at 37°C for 30 min. Slides were
counterstained with nuclear fast red. Specific controls included slides
in which the antibody was omitted (negative control), BAL cells treated with 500 µg/ml ribonuclease (RNase) for 30 min at 37°C before in
situ RT-PCR (negative control), and BAL cells in which GAPDH-specific primers were used for in situ RT-PCR (positive control; data not shown).
Immunocytochemistry.
Cytospin preparations of BAL cells collected 4 h after intratracheal
instillation of saline or LPS were dried and kept frozen at
70°C until processing. Slides with cells from control and LPS-treated rats were processed in parallel to avoid variations in
staining intensities attributable to processing. The
immunocytochemistry procedure and characterization of the SP antibody
have been described previously (6) with some modifications. After
fixation in cold picric acid-formaldehyde for 10 min and a rinse for 1 h in PBS with 0.3% Triton X-100 (PBS-TX), the cells were sequentially
incubated for 30 min at 30°C in ~100 µl of primary antibody
produced in rabbit against SP (Peninsula Laboratory, Belmont, CA)
diluted 1:200, then in biotin-labeled goat anti-rabbit immunoglobulin G
(Vector Laboratories, Burlingame, CA) diluted 1:100, and finally in
fluorescein-labeled avidin (Vector Labs) diluted 1:100, with intervening washes. PBS-TX was used for all washes and the diluents. Controls were processed through the same steps, substituting PBS-TX for
the primary antibody. The cells were viewed in a Zeiss fluorescence microscope, and digital images were captured using a cooled
charge-coupled device camera. The images were not processed further
except for the application of pseudocolor to mimic fluorescein.
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RESULTS |
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We initially detected PPT-I mRNA in rat AM using RT-PCR. Identity of the 212-bp PCR product was confirmed by Southern blotting, hybridization with a radiolabeled PPT-I-specific probe, and autoradiography (Fig. 1). Similarly, a band of the same size was detected in cDNA obtained from rat brain (positive control; Fig. 1). No PPT-I signal was detected in the no-template reaction or from peritoneal macrophages whether or not the cells had been stimulated with LPS (negative controls; Fig. 1). Appearance of the expected 440-bp GAPDH fragment from brain, AM, and peritoneal macrophages attested to the presence of amplified mRNA (positive controls; Fig. 1).
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Identity of the PPT-I cDNA was confirmed by sequence analysis of a purified 125-bp insert obtained by nested PCR. The sequence of the PCR product was identical to a previously reported sequence (17), confirming its identity as PPT-I (data not shown) (15). When this cDNA was radiolabeled and used as a probe for Northern analysis, a single band of ~1.3 kb in AM and brain (positive control; Fig. 2) was evident; however, no PPT-I mRNA was detected from rat peritoneal macrophages whether they were cultured as adherent or nonadherent cells (negative control). Expression of the proinflammatory chemokine MIP-2 mRNA was significantly induced in the same rat peritoneal macrophages by incubation with 10 µg/ml LPS for 4 h.
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Having shown that rat AM express PPT-I mRNA using RT-PCR, we used
Northern analysis to quantitate changes in mRNA levels in these cells
upon in vitro treatment with dexamethasone and/or LPS in three
separate cohorts of rats. Expression of PPT-I mRNA was induced after
incubation of AM with 10 µg/ml LPS, and this induction could be
markedly decreased by pretreatment with
107 M dexamethasone (Fig.
3). These experiments were repeated in vivo
with separate rat cohorts to determine whether PPT-I expression in BAL
cells was upregulated in a rat model of pulmonary inflammation and
downregulated by dexamethasone pretreatment. Acute lung inflammation was induced in vivo by 80 µg/kg LPS intratracheally and resulted in
an influx of neutrophils into the BAL fluid. BAL performed 4 h after
LPS treatment contained 69.8 ± 3.3% neutrophils compared with 1.0 ± 0.6% neutrophils in control animals. Similar to the in vitro
studies, PPT-I mRNA expression was significantly increased in BAL cells
from LPS-treated animals (Fig. 4). When the
hybridization to PPT-I mRNA was quantitated by scanning densitometry
and normalized to the GAPDH mRNA signal, PPT-I expression was enhanced
two- to fourfold. Similar to results in vitro, induction was eliminated by pretreatment of animals with 0.5 mg/kg dexamethasone (Fig. 4).
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We used in situ RT-PCR to further confirm expression of PPT-I mRNA and upregulation with LPS treatment. We also wished to determine whether other cells obtained by BAL, specifically neutrophils that constituted the majority of the cells recovered by BAL after LPS treatment, might contain PPT-I message. In situ RT-PCR of lung cells obtained by BAL using PPT-I-specific primers demonstrated a positive blue-black cytoplasmic precipitate in AM obtained from control animals (Fig. 5A). The cytoplasmic precipitate appeared darker and enhanced in animals treated with 80 µg/kg of LPS intratracheally 4 h before BAL (Fig. 5B). In addition, neutrophils that influxed into the lungs after LPS treatment also appeared to contain a positive blue-black cytoplasmic precipitate. No staining was detected in BAL cells treated with RNase or in cells in which the digoxigenin antibody was omitted (negative controls; Fig. 5, C and D, respectively). We further confirmed the in situ RT-PCR detection of PPT-I mRNA in BAL neutrophils obtained after intratracheal LPS treatment with Northern blotting. Neutrophils were purified from other cells in the BAL fluid, (>85% neutrophils), and Northern analysis was performed with 3 µg of total RNA per lane. A single band of 1.3 kb was detected in neutrophils, similar to that detected in AM (Fig. 6).
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Lung macrophages originate from chemotactic attraction of bone marrow-derived blood monocytes and local replication in the lung (19). Therefore, we purified blood mononuclear cells and used Northern analysis to determine whether these peripheral blood cells also express PPT-I mRNA. No signal was observed in rat blood mononuclear cells, although hybridizable GAPDH mRNA was readily detected from the same cells (Fig. 6).
We used immunocytochemistry to determine whether expression of PPT-I mRNA in AM was accompanied by protein expression. SP-like immunoreactivity was evident in macrophages from both control and LPS-treated groups (Fig. 7, A and B, respectively), but the fluorescence intensity was increased after LPS. Neutrophils observed in BAL after LPS treatment also appeared to be immunoreactive for SP. No cellular labeling was observed when PBS-TX was substituted for the primary antibody.
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DISCUSSION |
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These studies demonstrated that freshly lavaged rat AM expressed a single 1.3-kb transcript that was identical in size to PPT-I mRNA found in rat brain (Figs. 1 and 2). Expression of PPT-I mRNA was inducible with either in vitro or in vivo treatment with LPS and was downregulated by pretreatment with the synthetic glucocorticoid hormone dexamethasone (Figs. 3 and 4). AM subjected to in situ RT-PCR and hybridized with PPT-I-specific probes contained dark blue-black cytoplasmic precipitates in AM, consistent with a previous report demonstrating that PPT-I mRNA is located in the cytoplasm of cells (5) (Fig. 5). Immunocytochemical data paralleled the expression of PPT-I mRNA. Specifically, SP-like immunoreactivity was present in AM from both control and LPS-treated groups but the fluorescence intensity was increased after LPS (Fig. 7). Therefore, these studies suggested that, in addition to sensory C fibers, AM might be an unrecognized source of tachykinins in the lungs.
The in situ RT-PCR experiments indicated that neutrophils that influx into the lungs after in vivo LPS instillation also express PPT-I mRNA. Positive dark blue-black cytoplasmic precipitates were detected in neutrophils obtained by BAL that were similar to the cytoplasmic precipitate detected in AM (Fig. 5). When neutrophils were purified on a Nycodenz gradient from other BAL cells and Northern analysis performed, a single 1.3-kb size transcript was detected (Fig. 6). Neutrophils were observed in BAL after LPS treatment and appeared to be immunoreactive for SP, further supporting the conclusion that LPS-induced neutrophils expressed PPT-I mRNA and may be an additional source of SP during acute lung inflammation (Fig. 7). Rat peripheral mononuclear cells and peritoneal macrophages were negative for PPT-I mRNA, suggesting that PPT-I expression is specific to the AM and is not a general phenomenon of mononuclear phagocytes.
Multiple tachykinin peptides with related biological activities can be
derived from the PPT-I gene (5). The precursor RNA produced from the
PPT-I gene in rat is alternatively spliced to yield at least three
mRNAs that differ in exon usage within the protein coding region and
that are designated -,
-, and
-PPT-I (5, 17). Primers used in
these experiments corresponded to sequences that span exons 2 and 3 that are common to all of the known PPT-I mRNA splicing variants (Table
1). Primers were also designed to correspond to different exons so that
the PCR products studied would be derived from cDNA rather than from a
genomic DNA fragment, from sequences unlikely to undergo self and
mutual complementation, and from sequences with ~50% guanine and
cytosine content. It was not our intent to quantify
expression using RT-PCR and Southern blotting but rather to demonstrate
the presence or absence of PPT-I message in rat AM, brain (positive
control), and peritoneal macrophages (negative control). Northern
analysis was used for quantification of PPT-I message.
LPS, a potent activator of the immune system, the glucocorticoid
dexamethasone, and cytokines such as IL-1, IL-6, and tumor necrosis
factor- (TNF-
) have been shown to regulate PPT-I mRNA expression
in neuronal systems in vitro (10, 14, 27). Exposure of macrophages to
LPS leads to activation of the inducible transcription factor nuclear
factor-
B (NF-
B) (23). Transcriptional activation by NF-
B in
macrophages after LPS exposure has been reported after treatment with a
variety of immunomodulatory genes, including TNF-
, IL-1, IL-2, and
IL-6. Therefore, we hypothesized that LPS exposure might also increase
transcription of the PPT-I gene in AM similar to PPT-I upregulation
detected in the nervous system. We used a dose of LPS and a time course
that we had used previously to induce cytokines in AM in vitro and to
elicit acute pulmonary inflammation in vivo (7, 12). Our data indicated
that expression of PPT-I mRNA can be increased by in vivo exposure of
AM to LPS or by culturing AM in vitro with LPS, suggesting that, upon
activation, the ability of AM to make SP may be enhanced. When acute
lung inflammation was induced by in vivo treatment of rats with 80 µg/kg LPS intratracheally, PPT-I expression was induced two- to fourfold. A similar magnitude of induction of PPT-I mRNA is detected in
rat superior cervical ganglia after in vivo axotomy (25).
Dexamethasone treatment significantly inhibited PPT-I mRNA expression
in AM in vitro and virtually eliminated PPT-I message in animals
pretreated with dexamethasone before LPS instillation in vivo (Figs. 3
and 4). The mechanism(s) of dexamethasone-induced inhibition of PPT-I
expression is unknown, but previous reports have described repression
by in vitro treatment of neurons with dexamethasone (10). As with many
transcriptional regulators, dexamethasone can either stimulate or
inhibit gene expression. The DNA sequence element to which the occupied
dexamethasone receptor binds when mediating its negative effect, termed
the negative glucocorticoid response element (GRE), is distinct from
the GRE to which it binds when inducing gene expression. Although
putative transcriptional regulatory regions with homology to an
estrogen response element, adenosine 3',5'-cyclic
monophosphate response element, and serum response element have been
described (5, 14), the presence of GRE has not been demonstrated in the
promoter region of the PPT-I gene. Alternatively, dexamethasone might
inhibit PPT-I expression of LPS-stimulated macrophages by
interfering with the binding of the transcription factor NF-B to its
cis-element, as described for
dexamethasone suppression of IL-8 transcription by IL-1 (22).
Our data indicate that rat peritoneal macrophages do not synthesize PPT-I mRNA. We were unable to detect the PPT-I transcript using Northern analysis even when peritoneal macrophages were cultured as adherent cells or when activated by LPS treatment. In addition to PPT-I and GAPDH mRNA, we examined the expression of MIP-2 in the same peritoneal macrophages. MIP-2 is a potent neutrophil chemotactic and activating factor that functions during acute inflammation in vivo and is induced by LPS in vitro (12). MIP-2 expression in rat peritoneal macrophages was significantly induced by incubation with 10 µg/ml LPS for 4 h (data not shown) despite the absence of PPT-I mRNA. When peritoneal macrophage RNA was subjected to RT-PCR, which, in general, is ~1,000-10,000 times more sensitive than Northern analysis (30), we were still unable to detect a PPT-I signal. Stimulation of peritoneal macrophages in vitro with 10 µg/ml LPS followed by RT-PCR testing resulted in a very faint band, confirmed by Southern analysis, but was essentially undetected compared with the level found in rat brain and AM (Fig. 1). Thus we concluded that, even upon activation, sodium thioglycolate-elicited peritoneal macrophages contain little, if any, PPT-I mRNA message. These data are in contradiction to a previous study in which rat peritoneal macrophages were elicited by Freund's adjuvant, allowed to adhere to plastic petri dishes, and subsequently shown to contain PPT-I mRNA (3). The differences in these two studies most likely represent differences in the methods used to elicit peritoneal macrophages. Because both Freund's adjuvant and adherence to plastic "activates" macrophages, Bost et al. (3) were unable to determine whether constitutive expression of PPT-I occurs, but they hypothesized that unactivated macrophages would express little, if any, PPT-I mRNA. Nonetheless, their study suggests that, under specific conditions of activation, peritoneal macrophages may indeed express PPT-I mRNA.
We collected rat blood by intracardiac puncture and purified the mononuclear cells to address the question of whether PPT-I mRNA is present in circulating mononuclear cells or whether it is present after migration and final differentiation in the alveolar spaces. Based on the fact that we had been unable to detect a significant PPT-I signal from peritoneal macrophages, it was not surprising that blood mononuclear cells were negative for PPT-I.
The AM are the only macrophages in the body that are exposed to air (19). AM are theorized to be important targets for oxidative stress because of their role in pulmonary defense and their enhanced capacity for reactive O2 species production. Furthermore, reactive O2 species are known to activate gene expression through activation of transcription factors and/or modulation of the intracellular reduction-oxidation state of nuclear proteins (28). Work in our laboratory supports the hypothesis that oxidant-induced changes in the activity of regulatory proteins may influence cytokines by initiating transcription and/or by stabilizing mRNA (28). Oxidant-induced upregulation of mRNA levels may influence numerous genes and could account for expression of PPT-I mRNA in AM but not in peritoneal macrophages or peripheral mononuclear cells.
Our Northern analyses, RT-PCR, and in situ RT-PCR data indicated that AM and neutrophils recruited to the lung contained PPT-I mRNA and expressed SP. Immunocytochemical examination of AM suggested the presence of SP-like immunoreactivity that was increased after LPS, which paralleled PPT-I mRNA expression. Neutrophils were detected in the BAL after LPS treatment and appeared also to be immunoreactive for SP. We also attempted to measure the SP content of the AM and cell culture supernatant using an enzyme-linked immunosorbent assay. We have used this assay previously to measure SP content of the trachea or lungs (15). In the presence of a variety of peptidase inhibitors, we still found that the extraction of SP resulted in highly variable inconsistent results (data not shown). We suspect that we were unable to adequately inhibit all of the degradative enzymes present in the AM homogenate, resulting in enzymatic breakdown of the SP. Thus it is still unknown if AM release substantial quantities of SP and if tachykinins contribute to the inflammatory process. Despite evidence of PPT-I expression, similar attempts to extract and quantitate SP levels in eosinophils have also led to inconsistent results, making a quantitative interpretation difficult (21).
SP has been implicated as a contributing factor in both asthma and chronic bronchitis (29, 31). We have recently demonstrated that SP content in airways is significantly elevated in a rat model of chronic bronchitis (15). Nonetheless, the role of SP in AM and its possible contribution to inflammatory lung disease awaits further investigation. Our data suggest that immune cells in the lungs normally produce tachykinins and can increase this production during acute inflammation. SP-mediated complex bidirectional interactions between the immune (macrophages and neutrophils) and nervous systems (sensory C fibers) could significantly affect cellular proliferation, differentiation, and function.
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ACKNOWLEDGEMENTS |
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We thank Roger Morey, Dale Youngkin, Marshall Katler, and Thomas Donaghey for technical assistance.
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FOOTNOTES |
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This work was supported by National Institutes of Health Grants HL-19170, ES-05947, and ES-00002. C. R. Killingsworth is a Parker B. Francis Fellow in Pulmonary Research.
Address for reprint requests: C. R. Killingsworth, Univ. of Alabama at Birmingham, B147 Volker Hall, Box 201, 1670 Univ. Blvd., Birmingham, AL 35294-0019.
Received 25 March 1997; accepted in final form 14 August 1997.
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