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Physiol. Genomics 21: 230-242, 2005; doi:10.1152/physiolgenomics.00291.2004 Free Article
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Received 8 December 2004; accepted in final form 21 January 2005.
Physiological Genomics 21:230-242 (2005)
1094-8341/05 $8.00 © 2005 American Physiological Society

Inactivation of CD11b in a mouse transgenic model protects against sepsis-induced lung PMN infiltration and vascular injury

Xiao-Pei Gao *, Qinghui Liu *, Michael Broman , Dan Predescu , Randall S. Frey and Asrar B. Malik

Department of Pharmacology, The University of Illinois College of Medicine, Chicago, Illinois 60612


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
To inactivate chronically the ß2-integrin CD11b (Mac-1), we made a transgenic model in mice in which we expressed the CD11b antagonist polypeptide neutrophil inhibitory factor (NIF). Using these mice, we determined the in vivo effects of CD11b inactivation on polymorphonuclear leukocyte (PMN) function and acute lung injury (ALI) induced by Escherichia coli septicemia. In wild-type PMNs, CD11b expression was induced within 1 h after E. coli challenge, whereas this response was significantly reduced in NIF+/+ PMNs. Coimmunoprecipitation studies showed that NIF associated with CD11b in NIF+/+ PMNs. To validate the effectiveness of CD11b blockade, we compared PMN function in NIF+/+ and Mac-1-deficient (Mac-1–/–) mice. Adhesion of both Mac-1–/– and NIF+/+ PMNs to endothelial cells in response to LPS was reduced in both types of PMNs and fully blocked only by the addition of anti-CD11a monoclonal antibody. This finding is indicative of intact CD11a function in the NIF+/+ PMNs but the blockade of CD11b function. CD11b inactivation in NIF+/+ mice interfered with lung PMN infiltration induced by E. coli and prevented the increase in lung microvessel permeability and edema formation, with most of the protection seen in the 1-h period after the E. coli. Thus our results demonstrate that CD11b plays a crucial role in mediating lung PMN sequestration and vascular injury in the early phase of gram-negative septicemia. The NIF+/+ mouse model, in which CD11b is inactivated by binding to NIF, is a potentially useful model for in vivo assessment of the role of PMN CD11b in the mechanism of vascular inflammation.

neutrophil inhibitory factor; NIF+/+ mice; CD11b/CD18 integrin; neutrophil function; sepsis-induced lung injury; polymorphonuclear leukocyte


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
RECRUITMENT OF polymorphonuclear neutrophils (PMNs) in lungs has been implicated in the development of acute lung injury (ALI) (17, 28). The process of recruitment involves PMN adhesion to endothelial cells via ß2-integrins and transendothelial migration of PMNs into tissues (33). The PMN adhesion cascade comprises elements of tethering, rolling, activation, and firm adhesion (4, 33). Rolling and capture of PMNs are in large part mediated by endothelial P-selectin binding to P-selectin glycoprotein ligand-1 (PSGL-1). Stable adhesion of PMNs is mediated by the binding of PMN ß2-integrins CD11a/CD18 and CD11b/CD18 to intercellular adhesion molecule-1 (ICAM-1) on the endothelial cell surface (33). The ß2-integrins, lymphocyte function associated antigen-1 (LFA-1; CD11a/CD18) and Mac-1 (CD11b/CD18), are the major negative regulators of PMN rolling velocity; hence, deletion of ß2-integrin in mice (CD18–/– mice) was shown to increase the PMN rolling velocity (8, 16). ß2-Integrins are also capable of binding to multiple ligands; Mac-1 recognizes extracellular matrix proteins (fibronectin, laminin, collagen, and elastin), ICAM-1 and ICAM-2 (11), blood coagulation proteins [fibrinogen (1), factor X (2), and kininogen (15)], neutrophil inhibitory factor (NIF) (25), complement pathway product (iC3b) (3), and nonprotein ligands [e.g., lipopolysaccharide (LPS), ß-glucan (35), and heparin (6)]. The broad binding specificity of Mac-1 is not shared by LFA-1 even though the two have overlapping functional properties (44). Functional defects have been reported for Mac-1 gene deletion in mice; these include impaired PMN respiratory burst response (5), absence of PMN spreading and adhesion (5, 7, 21, 34), and reduced susceptibility to ischemia/reperfusion-induced tissue injury (32).

NIF is a 47-kDa protein isolated from the parasite Ancylostoma caninum that interferes with the host-defense response by apparently inactivating CD11b (25). Recombinant NIF inhibited the adhesion of activated human PMNs to the endothelium and release of PMN H2O2 over a concentration range (IC50 10–20 nM) (25). NIF binding sites on PMNs showed selective, high-affinity binding to the I-domain of CD11b (25) and low-affinity binding to CD11a in phorbol ester-stimulated PMNs (20). NIF protein has 257 amino acids preceded by a 17-amino acid leader sequence (25). The mature protein has 10 cysteines and 7 potential N-linked glycosylation sites (25). Mutagenesis analysis of CD11b identified Asp149, Arg151, Gly207, Tyr252, and Glu258 as being required for NIF binding (37, 45). Our studies using intravenous liposomes for delivery of NIF cDNA in mice showed that NIF expression in the lung interfered with PMN sequestration induced by sepsis (42, 43). However, it was not clear whether these results were the consequence of CD11b blockade per se, since the NIF expression levels were transient in these mice. Moreover, little is known about the consequences of chronic CD11b blockade induced by NIF expression on PMN function in vivo and its ability to interfere with lung PMN sequestration and vascular injury. To address the in vivo effects of persistent CD11b blockade, we developed a transgenic NIF+/+ mouse model.

Previous studies using genetic and immunopharmacological approaches have demonstrated that ß2-integrins not only participate in PMN adhesion but also regulate the PMN respiratory burst response (5, 7, 12, 23, 25). However, how chronic inactivation of CD11b interferes with PMN function and progression of lung microvascular injury induced by gram-negative septicemia has not been addressed. Therefore, we took advantage of the ability of NIF to bind to and inactivate CD11b in the transgenic NIF mice to address the effects of CD11b inactivation on the response to gram-negative septicemia.

In NIF+/+ mice, which constitutively express NIF mRNA and protein in all tissues, we observed that NIF protein was released into the circulation and circulating PMN CD11b function was impaired. The targeted CD11b inactivation induced in this manner interfered with PMN infiltration into lung tissue and prevented the E. coli-mediated increase in lung microvessel permeability and edema formation, in particular the responses occurring at 1 h after bacterial challenge. Thus NIF+/+ mice in which CD11b is specifically inactivated are a potentially useful model for assessment of the in vivo role of PMN CD11b integrin in the mechanism of vascular inflammation. Our results in NIF+/+ mice demonstrate the critical importance of CD11b in the mechanism of PMN sequestration and ALI induced by gram-negative sepsis.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals.
Transgenic mice were generated by introducing CMVnif DNA (5.7 kb) fragment into pronuclei of fertilized mouse embryos (CD1) as described (36). Southern blot hybridization and PCR were performed to identify the founder lines and transgenic offsprings. Mice homozygous for the transgene (NIF+/+) generated through breeding were used in all experiments. Wild-type CD1 (Wt) mice were used as controls. CD11b-deficient mice (Mac-1–/–) were obtained from Dr. Christie Ballantyne, Baylor College of Medicine, Houston, TX (21). All experimental procedures complied with institutional and National Institutes of Health guidelines for animal use, and approvals were obtained from the Institutional Review Board.

PMN surface ß2-integrin expression.
PMN surface expression of ß2-integrin was measured in whole blood using flow cytometry (19, 38). Briefly, blood from the right ventricle of control and genetic mice was drawn into a 1-ml syringe through a sterile 22-gauge needle at 1 h after intraperitoneal injection of 1 x 108 live E. coli (ATCC 25922). Peripheral blood samples from the individual mice were drawn into heparin-rinsed (10 U/ml) syringes and transferred into separate polypropylene tubes. Surface adhesion protein labeling was performed by incubation with phycoerythrin (PE)-conjugated anti-CD11a [monoclonal antibody (mAb) M17/4; Ebioscience] or PE-conjugated anti-CD11b (mAb M1/70; Ebioscience) antibodies. Isotype-matched control antibodies were used for each labeling (rat IgG2a-PE or rat IgG2b-PE; Pharmingen). Fluorescein isothiocyanate (FITC)-coupled anti-Ly-6G (BD Transduction Lab) was used as a counterstain for 1 h at 4°C. Then, 2.0 ml of lysing solution [fluorescence-activated cell sorting (FACS) lysing solution; Becton Dickinson, San Jose, CA] were added to each sample to hemolyze red blood cells. After centrifugation and washing with Dulbecco’s phosphate-buffered solution (D-PBS; Invitrogen, Carlsbad, CA), the remaining leukocytes were resuspended in 500 µl of 1% paraformaldehyde in D-PBS for fixation. The samples were then immediately analyzed by measuring PE fluorescence from the gated leukocyte population, using a Coulter EPICS Elite ESP (Coulter, Miami, FL). The forward and side light scatter profiles as well as FITC fluorescence were used to gate for PMN population. Fluorescence parameters were collected using four-decade logarithmic amplification. As a negative control, cells were stained with secondary antibody alone.

Confocal imaging.
Freshly isolated PMNs (2 x 106) were challenged with 1 µg/ml LPS (Serotype 0111:B4; Sigma, St. Louis, MO) at 37°C for 2 h and centrifuged for 5 min at 300 rpm, using a cytospin (Thermo Shandon, Pittsburgh, PA). Cells were permeabilized by methanol (purge and trap grade) fixation for 7 min at 20°C and blocked with 1% BSA in PBS. Thereafter, cells were incubated sequentially with the indicated primary antibodies, for 1 h each, at room temperature. After being washed five times for 5 min in 0.1% BSA in D-PBS (D-PBS-BSA), appropriate secondary Abs conjugated with rhodamine or fluorescein isothiocyanate (Jackson Immunoresearch Laboratories, West Grove, PA) were applied together, for an additional 1 h at room temperature. Cells were extensively washed in D-PBS-BSA and mounted on glass slides with Prolong Antifade mounting media (Molecular Probes, Eugene, OR), and images were acquired with a Zeiss LSM 510 confocal microscope.

PMN adhesion to endothelial cells.
Mouse bone marrow neutrophils were prepared as described with some modifications (5). Briefly, PMNs were isolated from femurs and tibias flushed with Ca2+/Mg2+-free Hanks’ balanced salt solution (HBSS)-BSA. The obtained marrow was centrifuged at 300 g, 4°C for 10 min, and resuspended in 3 ml of HBSS. The suspension was subjected to a Percoll step gradient, the gradient was then centrifuged, and cells were removed from the neutrophil-enriched fraction. This procedure yielded >95% PMN purity and >95% viability, assessed by Trypan blue exclusion. Cells were washed with Ca2+/Mg2+-free HBSS (for calcein AM labeling). The assay for PMN adhesion to endothelial cells was performed as described (27). Mouse lung vascular endothelial cells (MLVECs) were isolated (9) and grown to confluence in 96-well gelatin-coated plates. Bone marrow PMNs loaded with calcein AM (Molecular Probes) at 2 µg/ml for 30 min at room temperature were added to MLVECs pretreated with LPS (1 µg/ml) for 4 h at 37°C. We assessed PMN adhesion after treatment of PMNs with anti-CD11b mAb (M1/70) or anti-CD11a mAb (M17/4), each at a concentration of 10 µg/ml (BD Biosciences, San Diego, CA). The fluorescence readings were obtained with the PTI spectrofluorometer (Photon Technology International, Monmouth Junction, NJ) with detection at 485 and 535 nm, respectively. The percentage of adherent PMNs was calculated, and all assays were performed in duplicate.

FACS analysis of PMN oxidant generation.
Oxidant generation was measured by flow cytometry in isolated PMNs recovered in bronchoalveolar lavage (BAL) fluid (26). Briefly, BAL fluid (BALF) collected from E. coli-challenged mice was centrifuged, and the pellet containing leukocytes were resuspended in D-PBS. Dihydrorhodamine-123 (DHR, 3 mM; Sigma Aldrich) was added to all tubes except reagent blank (which had PBS added instead) and incubated at 37°C for 20 min in a water bath. The reaction was stopped by placement on ice. The samples were then immediately analyzed by measuring fluorescence of 15,000 events from the gated populations of PMNs and other leukocytes, using a Coulter EPICS Elite ESP (Coulter) with the laser set at 530 nm.

H2O2 production in adherent PMN.
H2O2 determinations, using the Amplex Red hydrogen peroxide kit, was performed according to the manufacturer’s instructions (Molecular Probes) with minor modifications (24). This assay is based on the detection of H2O2 using 10-acetyl-3,7-dihydroxyphenoxazine (Amplex Red reagent), a sensitive and stable probe for H2O2. PMNs at (1 x 105)/100 µl HBSS were added to MLVEC monolayers treated with LPS (1 µg/ml) for 2 h and then stimulated with fMLP (0.1 µM) for 10 min at 37°C to attain maximal oxidant production, according to the manufacturer’s recommendations. Fluorescence measurements of resorufin were performed at 563-nm excitation and 587-nm emission. All assays were performed in duplicate.

Lung PMN sequestration.
Lung tissue PMN sequestration was assessed by determining myeloperoxidase (MPO) activity in lungs and by morphometrically quantifying PMN infiltration as described (12, 13).

PMN trafficking into airspace.
The trachea was cannulated, and BAL was performed using 1 ml of D-PBS with 1 mM EDTA. BALF was centrifuged for 5 min at 300 rpm, using a cytospin (Shandon, Pittsburgh, PA), and BAL cells were stained with HEMA3 (Fisher, Chicago, IL) (12). Total PMN counts were determined with the use of a grid hemacytometer. Differential cell counts were enumerated on cytospine-prepared slides. A total of 500 cells were counted in cross-section per sample, and the numbers of PMNs were calculated as total cell count times the percentage of PMNs in BAL samples.

Dermal PMN extravasation.
Dorsal air pouches were created with minor modifications as described (7, 14). The air pouches were formed by injection of air (5 ml subcutaneously) on day 0 and day 3. On day 5, mice were challenged intraperitoneally with 1 x 108 live E. coli. Control mice were injected intraperitoneally with an equal volume of saline. The chemokine macrophage inflammatory chemokine-2 (MIP-2; Chemicon) in 0.1 ml of D-PBS was injected into the air pouch 30 min before collection of the air pouch lavage fluid. At 1 or 6 h after E. coli challenge, leukocytes were collected by flushing the air pouch with 8 ml of D-PBS. Lavage fluid was centrifuged at 300 g for 5 min, the resulting pellet was resuspended in 1 ml of D-PBS, and cells in an aliquot were counted using a grid hemacytometer; 90–95% of the cells in the exudates of air pouch were PMNs by staining with HEMA3.

PMN depletion/repletion studies.
Depletion of circulating PMNs followed by repletion of the PMNs was carried out as described (9). Briefly, mice were made neutropenic by intraperitoneal administration of 150 µl of rabbit anti-mouse PMN serum (Intercell Technologies, Hopewell, NJ). Circulating PMNs were not detectable at 24 h after serum injection, at which time PMN repletion experiments were performed by infusion of PMNs (1 x 106 cells) into the jugular vein. PMNs used for transfusion were isolated from bone marrow of either Wt, NIF+/+, or Mac-1–/– mice as described above. The infused PMNs were permitted to circulate for 1 h, at which time the repleted mice were challenged intraperitoneally with E. coli, and lung MPO activity was measured as described (12). The MPO activity was used as an index of lung PMN sequestration in the PMN-repleted mice. Control groups received equal amounts of normal rabbit serum by intraperitoneal injection.

Pulmonary microvascular permeability and lung water determinations.
Pulmonary capillary filtration coefficient (Kf,c) and lung wet weight increases were monitored as described (12, 13) to quantify pulmonary microvascular permeability to liquid and edema formation.

Immunoprecipitation and Western blotting.
Total protein was measured by bicinchoninic acid (BCA) analysis (31) (Pierce, Rockford, IL). Total PMN lysates were prepared using the following lysis buffer: 50 mM Tris·HCl, pH 8.0, + 1% TX-100; to this a protease inhibitor cocktail (AEBSF, pepstatin A, E-64, bestatin, leupeptin, and aprotinin) from Sigma was added before every experiment. The whole lung lysates were prepared, using the lysis buffer (1.5% Triton, 0.1% SDS, 0.5% sodium deoxycholate) plus a protease inhibitor cocktail and 1 mM phenylmethylsulfonyl fluoride (PMSF), as described above. Polyacrylamide electrophoresis was performed with precast SDS-PAGE gels purchased from Invitrogen. To determine the association of NIF with CD11a or CD11b, the whole lung lysates were precleared with protein A/G-agarose beads and incubated with 3 µg/ml of the first antibody for 2 h at 4°C before the addition of 20 µl of protein A/G-agarose (incubation at 4°C overnight). Immunoprecipitates were subjected to SDS-PAGE and Western blot analysis. Nitrocellulose membranes were blocked with 5% nonfat dried milk for 1 h and incubated with primary Abs overnight at 4°C. After washing three times with wash buffer, the membrane was incubated with horseradish peroxidase (HRP)-conjugated second antibodies for 1 h at room temperature. Protein bands were detected by enhanced chemiluminescence.

Statistical analysis.
Data are expressed as means ± SE. Statistical analysis was performed with two-way analysis of variance and the Newman-Keuls test for multiple comparisons. The numbers of experiments in the different groups are given in the figures. The criterion for statistical significance was P < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Characterization of NIF+/+ mouse model.
The mouse model was generated by introducing NIF cDNA linked downstream to cytomegalovirus (CMV) promoter by microinjection into fertilized mouse embryos (CD1). Integration of NIF cDNA into mouse genome was verified with Southern blot analysis of the mouse genomic DNA (Fig. 1A), and genotyping of offsprings was carried out using PCR (data not shown). We observed expression of NIF mRNA by Northern blot analysis (Fig. 1B). Western blotting showed that the NIF protein was present in all organs tested (heart, lung, liver, and kidney) and released into the circulation (Fig. 1C). Hematological analysis of NIF+/+ mice showed increased circulating PMN count and decreased numbers of lymphocytes compared with control (Fig. 1D); consequently, total leukocyte count was almost identical. Figure 2 shows midcell confocal images of PMNs with CD11b (green) and NIF (red). Both were associated in NIF+/+ PMNs but not Wt PMNs, indicating that NIF was present in the cytosol and membrane. Challenge of NIF+/+ PMNs with LPS (1 µg/ml, 2 h) increased NIF and CD11b association (Fig. 2).



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  		Fig. 1. Characterization of neutrophil inhibitory factor (NIF)+/+ mice. A: genomic DNA from mouse tail biopsies was digested with restriction endonuclease BamH I and then subjected to electrophoresis followed by Southern blot hybridization, using a 527-bp NIF cDNA as probe. The NIF transgene gives rise to 0.78-kb and 4.56-kb bands by the digestion, while the parental plasmid pCMVnif generates the same smaller fragment (0.78 kb) and a different larger band (5.16 kb). Samples 1–4 are 4 transgenic offsprings from breeding. N, negative control of CD1 mouse (wild type; Wt); P, plasmid pCMVnif; CMV, cytomegalovirus. B: Northern blot analysis of NIF transgene expression. Total RNAs (shown as ethidium bromide-staining gel) extracted from mouse lung and heart were probed with NIF cDNA probe (527 bp). C: NIF protein expression can be detected by Western blot analysis from tissues at various levels of abundance from NIF+/+ mice but not from tissues of Wt mice (CD1). Equal amounts of total protein (100 µg) were loaded per lane. Results are representative of 3 independent experiments; 1 of 3 comparable experiments is shown. D: total peripheral leukocyte counts and differentials of Wt and NIF+/+mice. Blood was drawn by cardiac puncture. Total no. of leukocytes in 100 µl was determined by Advia 120 (Bayer, Carpentersville, IL). Note that the polymorphonuclear leukocyte (PMN) count in NIF+/+mice is significantly elevated, while other cell populations remained unchanged. WBC, white blood cell; lymph, lymphocyte; mono, monocyte. Results are means ± SE of 8 experiments. *Increase in the genetic group compared with Wt group (P < 0.05).

 


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  		Fig. 2. Colocalization of NIF (red) with integrin {alpha}-chain CD11b (green), as indicated by orange and yellow staining (right) in the basal condition and after 2 h of lipopolysaccharide (LPS) stimulation in Wt and NIF+/+ PMNs.

 
Specific NIF binding to CD11b.
Immunoprecipitation with anti-NIF antibody (26) was performed to assess the relative binding of NIF to CD11a and CD11b in NIF+/+ mouse lungs and PMNs. Western blotting was carried out using anti-CD11b mAb followed by immunoprecipitation. CD11b expression increased within 1–6 h after E. coli challenge relative to basal values, with a greater response at 6 than 1 h (Fig. 3A). CD11b was coimmunoprecipitated with NIF in both mouse lung tissue and isolated mouse PMNs (Fig. 3A). No such response was seen in Wt PMNs. NIF association with CD11b was greater, concomitant with the increased expression of CD11b after E. coli challenge (Fig. 3B). In contrast, experiments showed no association between NIF and CD11a before or after E. coli challenge (Fig. 3C).



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  		Fig. 3. CD11b is the in vivo ligand for NIF in mouse lungs and PMNs. A: interaction of CD11b with NIF in lung and PMNs in NIF+/+ mice was demonstrated by precipitation of CD11b with anti-NIF antibody. Results are representative of 3 independent experiments. B: NIF is also immunoprecipitated from cell lysate by anti-CD11b antibody. Results are representative of 3 independent experiments. C: IP with anti-CD11a antibody did not result in precipitation of NIF. IB, immunoblot. Equal amounts of protein (200 µg of PMN lysate, 2 mg of lung lysate) were added to the immunoprecipitation (IP) reaction. Results are representative of 3 independent experiments.

 
Relative expression of CD11a and CD11b in NIF+/+ PMNs.
We assessed by flow cytometry the cell surface expression of CD11b in circulating Wt and NIF+/+ PMNs obtained after E. coli challenge of mice. As shown in Fig. 4, Wt PMN showed a significant increase in surface expression of CD11b at 1 h after intraperitoneal E. coli challenge (Fig. 4A, P < 0.05). This response in Wt PMNs was prevented by anti-CD11b mAb treatment of mice. In contrast, NIF+/+ PMNs showed ~55% reduction in CD11b expression compared with Wt PMNs, and this was also completely blocked by anti-CD11b mAb (Fig. 4A, P < 0.05). In a positive control experiment, we observed the expected absence of CD11b expression in Mac-1–/– PMNs (Fig. 4A). The increase in CD11a expression after E. coli was significantly less than CD11b expression (P < 0.01), and there was no difference in the level of increases in CD11a expression between Wt and NIF+/+ PMNs after E. coli challenge of mice (Fig. 4B). These results show that PMN-expressed NIF in NIF+/+ mice significantly reduced the CD11b expression in response to in vivo E. coli challenge, such that the response can be fully blocked by anti-CD11b mAb treatment. Results also show the selectivity of NIF in interfering with PMN CD11b expression since, the E. coli-induced increase in PMN CD11a expression was unaffected in NIF+/+ mice.



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  		Fig. 4. Flow cytometric analysis of cell surface CD11b/CD18 and CD11a/CD18 in circulating PMNs of Wt and NIF+/+ mice. A: mean phycoerythrin (PE) fluorescence intensity in CD11b-labeled Wt PMNs compared with NIF+/+ or Mac-1–/– PMNs obtained after 1 h of live E. coli (1 x 108) challenge (ip). B: PMN expression of CD11a in both Wt and NIF+/+ mice after E. coli challenge. PMNs were gated by forward and light angle scatter. Nonspecific PE fluorescence was determined by incubation with isotype- and fluorochrome-matched control antibodies. Isotype- and fluorochrome-matched control antibodies demonstrated minimal nonspecific fluorescence (data not shown). The relative mean PE fluorescence represents the average fluorescence from the CD11b- or CD11a-labeled population of isolated PMNs. Values are means ± SE; n = 3 separate experiments. *Increase in CD11b expression compared with basal (P < 0.05). **Decrease from E. coli-treated group (P < 0.05). {dagger}Decrease in CD11b expression of NIF+/+ PMNs compared with Wt PMNs.

 
Impaired NIF+/+ PMN adhesion to endothelial cells.
To examine the effects of the expressed NIF in NIF+/+ PMNs on PMN adhesion to endothelial cells, we challenged the PMNs with LPS (1 µg/ml, 4 h). LPS resulted in increased adhesion of Wt PMNs to MLVEC, whereas the increase of adhesion of NIF+/+ PMNs was 70% less than in Wt PMNs and 40% less in Mac-1 null PMNs than Wt PMNs (Fig, 5A). We also determined the effects of anti-CD11a or anti-CD11b mAb on PMN adhesion. In Wt PMNs, the LPS-induced increase in PMN adhesion was reduced by 80% in PMNs pretreated with either anti-CD11a or CD11b mAb. Adhesion of NIF+/+ and Mac-1–/– PMNs to endothelial cells was completely blocked with anti-CD11a mAb (P < 0.05), whereas reductions in the adhesion of NIF+/+ and Mac-1–/– PMNs were not significant with anti-CD11b mAb (Fig. 5A). To further evaluate the adhesive properties of NIF+/+ PMNs, we examined adhesion of PMNs to a CD11b-dependent ligand, fibrinogen (41). As shown in Fig. 5B, NIF+/+ PMN adhesion to fibrinogen was reduced by ~75% compared with Wt PMNs, and adhesion of Mac-1–/– PMNs to fibrinogen was completely abolished (Fig. 5B, P < 0.05). These data demonstrate that PMN NIF expression interfered with PMN adhesion by blocking CD11b and that some CD11b remained functionally active in NIF+/+ PMNs, since the PMN adhesion to fibrinogen was reduced by 75% in NIF+/+ PMNs and fully in Mac-1–/– PMNs.



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  		Fig. 5. Adhesion of Wt, NIF+/+, and Mac-1–/– PMNs to endothelial cells and fibrinogen. A: 1 x 105 PMNs were cocultured with mouse lung vascular endothelial cells (MLVECs) and stimulated with LPS (1 µg/ml) at 37°C for 4 h without or with anti-CD11a mAb (M17/4) and anti-CD11b mAb (M1/70) in a CO2 incubator. Treatment time before LPS was 15 min for both anti-CD11b and anti-CD11a mAbs (10 µg/ml). B: PMNs were added to a fibrinogen (100 µg/ml)-coated plate without or with LPS (1 µg/ml) stimulation at 37 °C for 4 h in a CO2 incubator. Results are means ± SE of 3 independent experiments. *Decrease in PMN adhesion of Wt group compared with LPS alone (P < 0.05). {dagger}Decrease in PMN adhesion of genetic PMNs compared with Wt PMNs (P < 0.05). *{dagger}Decrease compared with LPS alone in the genetic group (P < 0.05).

 
Impaired lung PMN sequestration in NIF+/+ mice.
Lung tissue PMN sequestration (as measured morphometrically) increased after E. coli challenge in a time-dependent manner, with a threefold increase from basal level at 1 h (Fig. 6A). The value peaked at 1 h and decreased at 6 h (Fig. 6A). Increases in lung PMN sequestration were reduced in NIF+/+ by 60% compared with Wt mice at 1 h and by 25% at 6 h (Fig. 6A, P < 0.05). Lung tissue MPO activity (another assay of lung PMN sequestration) also increased after E. coli challenge in Wt mice, reaching a maximum at 1 h (Fig. 6B). Lung MPO activity in NIF+/+ mice was reduced by 80% at 1 h and by 30% at 6 h after E. coli compared with Wt (Fig. 6B, P < 0.05). Thus the results indicate that early 1-h PMN sequestration induced by E. coli challenge was reduced to a greater extent in NIF+/+ mice compared with the 6-h response, suggesting the greater involvement of CD11b in mediating the early lung PMN uptake response.



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  		Fig. 6. Morphometric analysis of lung PMN sequestration and lung myeloperoxidase (MPO) activity. Mice were challenged for 0.5, 1, or 6 h with 1 x 108 E. coli (ip), and lungs were harvested for morphometric assay of PMNs in lungs and MPO activity. A: time course of lung PMN sequestration in Wt and NIF+/+ mice. B: time course of tissue MPO activity in lungs of Wt and NIF+/+ mice after E. coli challenge. Results are means ± SE of 6 experiments. Significant changes are as follows. *Increase compared with basal (P < 0.05). {dagger}Decrease relative to Wt group (P < 0.05).

 
Contribution of NIF expression in PMNs to the reduction in lung PMN sequestration.
We addressed whether NIF expressed in PMNs was responsible for the observed reduction in lung PMN sequestration in response E. coli. Studies were made by making mice neutropenic and then infusing NIF+/+ PMNs; these mice were challenged with E. coli, and PMN sequestration was assessed at 1 h after challenge (see MATERIALS AND METHODS). Results showed that Wt mice with circulating NIF+/+ PMNs or Mac-1–/– PMNs had ~70% reduction in lung PMN uptake at 1 h after E. coli challenge in each case compared with the control group (Fig. 7, bar 2 vs. bars 5 and 6). These results demonstrate that the reduction in lung tissue PMN uptake at 1 h after E. coli challenge in NIF+/+ mice (Fig. 6) is the result of inhibition of CD11b in the circulating PMNs.



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  		Fig. 7. Lung tissue PMN influx in neutropenic mice after E. coli challenge. Neutrophil depletion was achieved by injection (ip) of anti-neutrophil Ab 24 h before 1 x 108 E. coli (see MATERIALS AND METHODS). Bars 1 and 2: control response of Wt mice before and 1 h after E. coli challenge. Bars 3–6: lung MPO activity in Wt mice that have been made neutropenic. Bars 7–10: lung MPO activity in neutropenic NIF+/+ mice that have been repleted with PMN isolated from Wt or genetic mice. Data are means ± SE of 6 animals/group. Statistically significant differences from the reference group are as follows. *Compared with no E. coli challenge (basal value, P < 0.05). *{dagger}Decrease with repletion of genetic PMNs compared with repletion of Wt PMNs into neutropenic mice post-E. coli challenge (P < 0.05). {dagger}Decreased MPO activity after repletion of Wt PMNs in the genetically modified group compared with the Wt group post-E. coli challenge (P < 0.05).

 
To address the possible contribution of release of NIF in interfering with PMN uptake, studies were also made in NIF+/+ mice infused with Wt PMNs. E. coli challenge of these mice resulted in ~40% reduction in lung PMN uptake compared with control (Fig. 7, bar 2 vs. bar 8), indicating that NIF release is capable of binding to and inhibiting PMN uptake in vivo. Infusion of NIF+/+ PMNs or Mac-1–/– PMNs in these NIF+/+ mice further decreased lung PMN uptake induced by E. coli to the levels seen with infusion of NIF+/+ PMNs or Mac-1–/– PMNs in Wt mice (Fig. 7, bars 5 and 6 vs. bars 9 and 10). These results suggest that the release of NIF into the circulation (Fig. 1) and its binding to PMNs (Fig. 4) contribute to blocking lung PMN uptake.

PMN transalveolar migration vs. dermal extravasation in NIF+/+ mice.
To address the role of CD11b in regulating PMN migration in lungs, Wt and NIF+/+ mice were challenged with E. coli, and PMNs were counted in BALF. E. coli caused a significant increase in BALF PMNs in Wt mice compared with the saline-treated control. There was a 50% reduction of transalveolar PMN migration in NIF+/+ and a 25% reduction in Mac-1–/– mice at 6 h compared with Wt mice (Fig. 8A, P < 0.05). Reductions of PMN migration into the airspace in NIF+/+ and Mac-1–/– mice indicate an important role of inactivation of CD11b in regulating the PMN migration in vivo sepsis model.



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  		Fig. 8. Transalveolar PMN migration and dermal PMN extravasation in vivo. A: PMN migration into airspace of Wt or CD11b-blockaded (NIF+/+ and Mac-1–/–) mice after 1 h of E. coli challenge (1 x 108 ip). PMN counts in bronchoalveolar lavage fluid (BALF) were determined post-E. coli (see MATERIALS AND METHODS). B and C: PMN extravation of Wt, NIF+/+, and Mac-1–/– in air pouch. Subcutaneous air pouch was prepared as described in MATERIALS AND METHODS. Total PMN counts were determined using a grid hemacytometer. Values are means ± SE; in each experimental group, at least 6 mice were used. *Increase compared with basal (P < 0.05). {dagger}Decrease relative to Wt group (P < 0.05).

 
To evaluate CD11b-dependent dermal leukocyte extravasation, we also examined PMN subcutaneous migration using the air pouch model in Wt, NIF+/+, and Mac-1–/– mice. We first challenged the mice with E. coli (ip) or saline, and the chemokine MIP-2 was then injected into the air pouch of mice at 30 min before the pouch lavage fluid was collected. As shown in Fig. 8B, at 1 h after E. coli challenge, MIP-2 caused a threefold increase in PMN migration in Wt mice compared with the saline control that persisted up to 6 h after E. coli. The migration of PMNs into the air pouch was unaffected in NIF+/+ and Mac-1–/– mice (Fig. 8, B and C). These results indicate that the migration of PMNs into the airspace is in part the result of CD11b, whereas PMN transmigration across the dermal vessel is CD11b independent.

Oxidant production in NIF+/+ PMNs.
Oxidant production by activated PMN was measured, using flow cytometry, in NIF+/+ and Wt mice. Oxidant production of PMNs obtained from BALF was increased at 1 h after E. coli challenge (Fig. 9A), whereas this response was significantly reduced in NIF+/+ mice (Fig. 9A, P < 0.05). Studies were also made by stimulating PMNs cocultured with mouse endothelial cells for 2 h with LPS, followed by challenge of the adherent PMNs with fMLP for 10 min. This resulted in a marked ninefold increase in H2O2 production (Fig. 9B, P < 0.05). The response was reduced 65% in NIF+/+ and 55% in Mac-1–/– PMNs (Fig. 9B, P < 0.05). These results indicate that E. coli challenge of mice resulted in PMN oxidant production, which was in part CD11b mediated, since the response was reduced in NIF+/+and Mac-1–/– PMNs.



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  		Fig. 9. Oxidant production in PMNs in vivo and PMNs adherent to endothelial cells. A: oxidant generation from PMNs in BALF (as measured by flow cytometry) collected from mice challenged with E. coli (1 x 108 ip). Oxidant generation of Wt PMNs reached maximum at 1 h, as shown by significant right shift. NIF+/+ PMNs had a reduced oxidant generation response, as shown by smaller right shift compared with Wt PMNs. Values are means ± SE; n = 3 separate experiments (insets). B: oxidant generation in adherent PMNs. H2O2 production by PMNs in PMN-endothelial coculture system was measured as described in MATERIALS AND METHODS. Cells (1 x 105) in the coculture were stimulated with LPS (1 µg/ml) at 37°C for 2 h and then with fMLP (0.1 µM, 10 min) in a CO2 incubator. *Compared with no LPS+fMLP stimulation (basal value, P < 0.05). {dagger}Decreased in genetic group compared with Wt group post-LPS+fMLP stimulation (P < 0.05). Results are means ± SE of 3 independent experiments.

 
Lung microvessel permeability and edema formation induced by E. coli challenge in CD11b-blockaded NIF+/+ mice.
E. coli challenge resulted in increases in Kf,c (vessel wall liquid permeability) and lung wet weight. Increases in Kf,c and lung wet weight were time dependent in Wt control mice (Fig. 10). Increases in Kf,c after E. coli challenge were reduced at 1 h by 70% in NIF+/+ mice and 60% in Mac-1–/– mice compared with Wt mice, and by 50% in NIF+/+ mice and 40% in Mac-1–/– mice at 6 h after E. coli challenge (Fig. 10A). Lung wet weight gain was also significantly reduced in NIF+/+ mice compared with Wt mice at both time points (Fig. 10B), indicating that expression of NIF protected the mice from sepsis-induced acute lung injury. The greater protection against lung vascular injury at 1 h after E. coli in NIF+/+ and Mac-1–/– mice than at 6 h is consistent with the greater contribution of CD11b in the mechanism of lung PMN uptake at 1 vs. 6 h (Fig. 6, A and B).



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  		Fig. 10. Lung microvessel permeability and edema formation in NIF+/+ mice. A: time course of microvessel liquid permeability change (measured as capillary filtration coefficient; Kf,c) was determined in lungs isolated 1 and 6 h after E. coli challenge (1 x 108 ip). Values are means ± SE; n = 6 independent experiments. *Compared with no E. coli (basal value). {dagger}Decrease in Kf,c value of genetic mice relative to Wt mice post-E. coli challenge (P < 0.05). B: time course of pulmonary edema formation (measured by increase in isogravimetric water content) in lungs isolated 1 and 6 h after E. coli challenge. *Compared with basal value (0 min). {dagger}Decrease in lung wet weights of genetic mice relative to Wt mice post-E. coli challenge (P < 0.05). Results are means ± SE of 6 independent experiments.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
We describe herein a novel transgenic NIF+/+ mouse model in which CD11b is chronically blocked by the constitutive release of NIF, the CD11b polypeptide antagonist (20, 25). Using this model, we compared the effects of gram-negative septicemia induced by E. coli challenge with CD11b deficiency in Mac-1–/– mice. Our results show that NIF protein expression significantly reduced PMN CD11b activity, which is typically upregulated in response to E. coli challenge. The effect of NIF in blocking CD11b was selective, since the increase in PMN CD11a expression induced by E. coli was unaffected. NIF expression in PMNs interfered with PMN adhesion by blocking CD11b function; however, it appears that some component of CD11b remained active in NIF+/+ PMNs, since adhesion of these PMNs to fibrinogen, which solely binds to CD11b (7), was reduced by 75%, whereas it was fully blocked in Mac-1–/– PMNs. We also observed that lung PMN sequestration occurring at 1 h after E. coli challenge was reduced to a greater extent in NIF+/+ mice compared with the 6-h response, indicating the greater involvement of CD11b in mediating the early lung PMN uptake response. Studies in Wt mice repleted with NIF+/+ PMNs demonstrated that the reduction in lung PMN uptake was largely attributable to inhibition of CD11b function of the PMNs. We also observed reduction of PMN migration into the airspace in both NIF+/+ and Mac-1–/– mice, indicating that PMN migration into the airspace significantly relies on CD11b, whereas PMN transmigration across the dermal vessel is fully CD11b independent. In addition to the effect of NIF in blocking CD11b function, PMN oxidant production induced by E. coli challenge of mice also required CD11b, since the response was reduced equally in both NIF+/+ and Mac-1–/– PMNs. Finally, a greater protection against lung vascular injury was seen at 1 h after E. coli in NIF+/+ and Mac-1–/– mice than at 6 h, indicative of a greater contribution of CD11b in mediating the early phase of lung PMN uptake.

PMNs are the first cells to transmigrate across the endothelial barrier to a site of infection in response to signals generated in the vasculature and tissues. PMNs and other leukocytes utilize selectins and ß2-integrins in a sequence of overlapping events to traverse postcapillary venules (18). PMN recruitment proceeds in a cascade-like fashion, from initial capture to decreased velocity to firm adhesion and to transmigration (33). In the present study, we addressed the effects of functionally disabling CD11b by transgenically expressing the inhibitory protein NIF. NIF cDNA contains a 5'-end sequence encoding a signal peptide that induces secretion of the synthesized NIF (25), as evident by the presence of the protein in the circulation. Expression of NIF protein induced the formation of the CD11b-NIF complex, resulting in the impairment of CD11b function. We showed by confocal imaging that the expressed NIF protein interacted with CD11b at the cell surface and cytosol, suggesting that NIF was capable of blocking both pools of CD11b. Our previous study showed that NIF can bind with low affinity to CD11a after activation of PMN with phorbol ester (phorbol 12-myristate 13-acetate; PMA) (20); however, in the present study, the effect of NIF in blocking CD11b function after sepsis was specific in that NIF did not bind to CD11a.

Previous studies have evaluated the contributions of CD11a, CD11b, and CD18 in mediating leukocyte extravasation using approaches such as deletion of these genes and mAbs directed against these integrins (21, 23). Studies showed that leukocyte migration across dermal vessels (in a model in which TNF-{alpha} was injected into a subcutaneous air pouch) was significantly reduced in CD11a- and CD18-null mice, but marked influx was seen in CD11b-null mice (7). These studies suggested that the absence of CD11b may have unmasked a compensatory role of CD11a in mediating PMN emigration across dermal vessels. Mizgerd et al. (23) demonstrated CD18-independent pathways as being responsible for PMN emigration during pneumonia or peritonitis, since PMN emigration at these sites persisted in CD18-null mice. Walzog et al. (40), however, observed a reduced number of extravasated leukocytes into the inflamed peritoneal cavity of CD18–/– mice compared with Wt mice, indicative of a role of CD18 in the mechanism of PMN migration in the peritoneum. CD11b-null mice (Mac-1–/–) showed normal circulating PMNs, and other blood cells isolated from these mice lost their ability to adhere to fibrinogen-coated glass and showed reduced aggregation (21). In a model of inflammation induced by injection of thioglycolate into the peritoneal cavity (21), the accumulation of PMNs in Mac-1–/– mice was comparable to that in Wt mice, but emigration of the Mac-1–/– PMNs was more dependent on CD11a than in Wt PMNs (i.e., anti-CD11a mAb inhibited 78% of PMN accumulation in Mac-1–/– mice vs. 58% inhibition in Wt mice). This finding suggested a compensatory effect from CD11a in CD11b-null mice, consistent with the overlapping roles of CD11a and CD11b integrins on leukocyte emigration from the vasculature (7). Given the often confounding results seen in these gene deletion models, perhaps secondary to compensations and the complexity of Mac-1 integrin binding specificity and its multiple functions, in the present study we undertook the development of a model of targeted CD11b inactivation (and hence the NIF+/+ mice). Using these mice, our intent was to address the role of CD11b in the mechanism of lung PMN sequestration and vascular injury.

A question arises as to whether the amount of NIF expressed in these mice is sufficient to block both CD11b expression and function of PMNs. We showed that adhesion of NIF+/+ PMNs to endothelial cells was reduced by 70%, and it was completely inhibited with the addition of anti-CD11a mAb, indicating that NIF blockade of CD11b significantly impaired PMN adhesion. We also evaluated the adhesive properties of NIF+/+ PMNs to the CD11b ligand fibrinogen (41). NIF+/+ PMN adhesion to fibrinogen was reduced by 75% compared with Wt PMNs, whereas adhesion of Mac-1–/– PMNs to fibrinogen was completely abolished. These data demonstrate that PMN NIF expression interfered with PMN adhesion by blocking CD11b but that a component of CD11b still remains functionally active in the NIF+/+ PMNs. We observed by flow cytometry that NIF expression in NIF+/+ PMNs also significantly reduced the CD11b expression in response to the E. coli challenge. This effect of NIF on PMN CD11b expression was selective, since the increase in PMN CD11a expression was unaffected in NIF+/+ PMNs. Thus the NIF+/+ mice represent a new model distinct from the CD11b-null model (Mac-1–/– mice), in that the latter has a complete defect in the expression of the protein, whereas NIF+/+ mice have normal CD11b protein expression but its function is chronically blocked.

Another important difference between the NIF+/+ mice and the previously described Mac-1–/– mice (7, 21) is the elevated basal circulating PMN count in the former group. This finding can be explained by impairment of PMN extravasation as the result of CD11b blockade in NIF+/+ mice. It is possible that the compensatory expression of CD11a in Mac-1–/– mice may have activated PMN transmigration (21), resulting in the relatively normal circulating PMN count in these mice.

We determined lung PMN sequestration in NIF+/+ mice in response to E. coli challenge. Lung tissue PMN sequestration measured morphometrically and by the MPO activity assay increased after E. coli challenge in a time-dependent manner, with a peak value seen at 1 h after bacterial challenge. The increase in lung PMN sequestration was reduced by 60–80% in NIF+/+ mice compared with Wt mice at 1 h and by 25–30% at 6 h after challenge. These results demonstrate a crucial role of CD11b in mediating the early phase of PMN sequestration, indicating the greater involvement of CD11b in the initial phase of PMN uptake than at later time points.

We addressed in PMN depletion/repletion studies whether NIF protein expressed in PMNs is per se responsible for the observed reduction in lung PMN sequestration after E. coli challenge. Mice were made neutropenic and then infused with NIF+/+ PMNs, followed by challenge with E. coli and assessment of PMN sequestration at 1 h after challenge. Interestingly, Wt mice with circulating NIF+/+ PMNs or Mac-1–/– PMNs had ~70% reduction in lung PMN uptake after E. coli challenge compared with the control group. Thus the reduction in lung tissue PMN uptake at 1 h after E. coli challenge in NIF+/+ mice appears to be largely due to the result of inhibition of CD11b function of the circulating PMNs.

It is also possible that NIF expressed in the tissues of NIF+/+ mice is another important source of NIF responsible for the PMN CD11b blockade. To address the contribution of NIF release from tissues in interfering with PMN uptake, studies were made in NIF+/+ mice transfused with Wt PMNs. E. coli challenge of these mice resulted in 40% reduction in lung PMN uptake compared with controls, indicating that NIF release is capable of partially reducing PMN uptake in vivo. Transfusion of NIF+/+ PMNs or Mac-1–/– PMNs in these NIF+/+ mice, however, decreased lung PMN uptake induced by E. coli to the levels seen with transfusion of NIF+/+ PMNs or Mac-1–/– PMNs into Wt mice. Together, these results demonstrate that the release of NIF into the circulation and its binding to circulating PMNs are only partly responsible for reducing lung PMN uptake. A much greater reduction in lung PMN uptake occurred when PMNs expressed the copy of the NIF transgene. Our findings are consistent with evidence that CD11b, unlike CD11a, is markedly upregulated on the PMN surface in response to inflammatory stimuli (7); thus these observations help to explain the effect of inactivation of PMN CD11b in NIF+/+ PMNs in significantly reducing the lung PMN sequestration.

To assess the relative contributions of CD11b in regulating transendothelial PMN trafficking in different vascular beds, we compared PMN extravasation into the airspace and across dermal microvessels. We observed a 50% reduction of transalveolar PMN migration in NIF+/+ mice and a 25% reduction in Mac-1–/– mice at 6 h after E. coli compared with Wt mice, thus pointing to an important role of NIF inactivation of CD11b in interfering with PMN migration across the lung capillary-alveolar barrier. To assess CD11b dependence of dermal leukocyte extravasation, we examined subcutaneous migration of PMNs using the air pouch model. The migration of PMNs into the air pouch was unaffected in NIF+/+ and Mac-1–/– mice, indicating that PMN transmigration across dermal vessels is CD11b independent. Frevert at al. (10) also showed that different mechanisms are utilized in lungs vs. skin in signaling PMN emigration, in that the response in the skin was less IL-8 dependent than in lungs. Thus CD11b is a key determinant of accumulation of PMNs within the pulmonary microvascular bed and migration into the airspace induced by gram-negative septicemia.

Oxidant generation by PMNs is a distinguishing feature of activation of PMNs. This response, stimulated by chemotactic factors, was markedly augmented by CD11b-dependent adhesion via the interaction of CD11b with ICAM-1 expressed on the endothelial cell surface (22, 29). CD11b was also shown to be associated with components of the NADPH complex, p47phox and gp91phox, after TNF-{alpha} challenge (unpublished data). We observed a reduction in the generation of oxidants in transmigrated NIF+/+ PMNs obtained by bronchoalveolar lavage after E. coli challenge as well as adherent PMNs after LPS stimulation compared with Wt PMNs. This finding indicates that inactivation of CD11b in NIF+/+ PMNs blocked the pathways responsible for production of oxidants. The results are in agreement with studies showing an important role of CD11b in mediating PMN oxidant production (30, 39). Thus, on the basis of these data obtained using NIF+/+ PMNs, it appears that CD11b not only regulates lung PMN sequestration induced by gram-negative sepsis but also PMN oxidant production.

Because endothelial injury induced by PMNs adherent to endothelial cells is an important factor in the pathogenesis of ALI, we challenged mice with E. coli and quantified the lung microvessel filtration coefficient (Kf,c), a measure of vascular permeability, and the gain in lung wet weight, a measure of tissue edema formation. The increase of lung vascular permeability induced by sepsis was significantly reduced in NIF+/+ mice at both the 1- and 6-h points after E. coli challenge, but the protection was greater at the earlier time, paralleling the marked reduction in the initial lung tissue PMN sequestration. Thus we conclude that CD11b engagement contributes to the mechanism of the early phase of lung PMN infiltration and resultant vascular injury induced by gram-negative sepsis.


   GRANTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This study was supported by National Heart, Lung, and Blood Institute Grants HL-64573, HL-45638, and HL-46350.


   ACKNOWLEDGMENTS
 
We acknowledge Drs. A. Rahman and N. Xu for making the CMV-NIF cDNA construct. We also thank Dr. K. Ley for critical reading, Dr. C. Ballantyne for supplying the Mac-1-deficient mice, and the Flow Cytometry Facility at the Research Resources Center of the University of Illinois at Chicago for technical assistance.


   FOOTNOTES
 
Article published online before print. See web site for date of publication (http://physiolgenomics.physiology.org).

Address for reprint requests and other correspondence: A. Malik, Dept. of Pharmacology, College of Medicine, The Univ. of Illinois, 835 South Wolcott Ave., Chicago, IL 60612-7343 (E-mail: abmalik{at}uic.edu).

10.1152/physiolgenomics.00291.2004.

* Xiao-Pei Gao and Qinghui Liu contributed equally to this work. Back


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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