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Published ahead of print on November 24, 2004, doi:10.1165/rcmb.2004-0108OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 32, pp. 108-117, 2005
© 2005 American Thoracic Society
DOI: 10.1165/rcmb.2004-0108OC

Monocytes Recruited to the Lungs of Mice during Immune Inflammation Ingest Apoptotic Cells Poorly

Jeffrey H. Jennings, Derek J. Linderman, Bin Hu, Joanne Sonstein and Jeffrey L. Curtis

Division of Pulmonary and Critical Care Medicine, Department of Internal Medicine; the Comprehensive Cancer Center, and the Graduate Program in Immunology, University of Michigan Health System; and the Pulmonary and Critical Care Medicine Section, Medical Service, Department of Veterans Affairs Medical Center, Ann Arbor, Michigan

Correspondence and requests for reprints should be addressed to Jeffrey L. Curtis, M.D., Pulmonary & Critical Care Medicine Section (506/111G), Department of Veterans Affairs Medical Center, 2215 Fuller Road; Ann Arbor, MI 48105-2303. E-mail: jlcurtis{at}umich.edu


   Abstract
Top
 Abstract
Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 
Apoptotic cells must be cleared to resolve inflammation, but few resident alveolar macrophages (AMø) from normal lungs ingest apoptotic cells. We examined how Mø ingestion of apoptotic cells is altered during immune inflammation induced by intratracheal challenge of primed C57BL/6 mice using sheep red blood cells. Resident AMø were labeled in situ before challenge using intravenous PKH26 to distinguish them from recruited monocytes. Using flow cytometry, we identified phagocytosis of fluorescently-labeled apoptotic thymocytes by alveolar mononuclear phagocytes in vitro and in vivo, and measured surface molecule expression. Intratracheal challenge induced rapid recruitment of monocytes, peaking at Day 3 and decreasing thereafter, whereas numbers of resident AMø did not change significantly. At all times, the percentage of phagocytes ingesting apoptotic thymocytes in vitro was greater among resident AMø (28–45%) than among recruited monocytes (9–19%), but was low in both cell types relative to ingestion of immunoglobulin-opsonized targets. There was also a nonsignificant trend toward lower ingestion by monocytes in vivo. MerTK, a receptor tyrosine kinase crucial for apoptotic cell phagocytosis, was expressed by resident AMø, but not by recruited monocytes. Relative to resident AMø, monocytes recruited to the alveolus ingest apoptotic cells meagerly, possibly due to absence of MerTK expression.

Key Words: apoptosis • adhesion molecules • mice, inbred strains • macrophage • receptor tyrosine kinase


   Introduction
Top
 Abstract
 Introduction
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 
Efficient clearance of apoptotic leukocytes is crucial for the resolution of inflammation (1). Defective clearance of apoptotic bodies is associated with autoimmunity in some mouse strains (2) and appears to be involved in the pathogenesis of systemic lupus erythematosus in humans (3). During resolving inflammation, apoptotic leukocytes are believed to be cleared predominantly by resident tissue macrophages (Mø), which are abundant in virtually all organs (4). Mø that ingest apoptotic cells accelerate the resolution of inflammation by producing anti-inflammatory mediators such as transforming growth factor-ß and prostaglandin E2 (5). Because these mediators also have the potential to induce fibrosis and to impair antimicrobial defenses, understanding the molecular mechanisms of apoptotic cell clearance by Mø is an important goal.

How apoptotic cells are cleared from the lungs remains incompletely defined. Decreased phagocytosis of apoptotic cells by resident alveolar Mø (AMø), relative to their own ingestion of other targets or to ingestion of apoptotic cells by other tissue Mø, has been demonstrated in rabbits, mice, and most recently in humans (6–8). In mice, the most thoroughly studied species, relatively deficient apoptotic cell phagocytosis was seen both in vitro in seven murine strains and in vivo, was specific for apoptotic cells (as opposed to three other phagocytic targets), and extended to ingestion of apoptotic neutrophils (7, 9). This deficiency is explained in part by reduced adhesion of apoptotic cells (10), and in part by marked reduction in the expression of protein kinase C (PKC) ßII, which is necessary for apoptotic cell ingestion (11). Nevertheless, AMø share many features of apoptotic cell recognition with the avidly phagocytic resident peritoneal Mø (PMø) (9), including reliance on the stereospecific phosphatidlyserine receptor (PSR) (11, 12) and the receptor tyrosine kinase MerTK (also known as Tyro12) (2, 10). This combination of unique and shared features implies that the alveolar surface constitutes an ideal environment in which to investigate the mechanisms and consequences of altered apoptotic cell clearance.

The reduced capacity of resident murine AMø to ingest apoptotic cells raises two questions. First, does phagocytosis of apoptotic cells by resident AMø increase during resolving inflammation, when the burden of cells to be cleared rises? Second, might apoptotic cells actually be cleared under these circumstances primarily by recruited monocytes? In vitro, human monocyte-derived macrophages can clearly develop the capacity to ingest apoptotic cells, and the rate at which they develop this capacity is accelerated by cytokines that are likely to be expressed during resolving inflammation (13). However, it is unproven whether monocytes actually develop this capacity in vivo, and if so, over what time period.

To address these questions experimentally, we analyzed phagocytosis of apoptotic thymocytes by murine lung mononuclear cell phagocytes in vitro and in vivo, distinguishing resident AMø from monocytes recruited to the alveolar spaces by a previously described vital staining technique (14). This technique, which relies on differential uptake of the vital dye PKH26 by phagocytic resident Mø versus the nonphagocytic peripheral blood monocyte, has been used extensively to analyze monocyte recruitment in murine models of lung inflammation (14–16). We employed a well characterized model of CD4 T cell–dependent lung inflammation, the secondary immune response of primed mice to intratracheal challenge with the particulate antigen sheep red blood cells (SRBC) (17). A salient feature of this experimental model system in addressing the current questions is that we have previously shown that a substantial fraction of alveolar lymphocytes are apoptotic throughout the course of this response (18). We found that the fraction of resident AMø ingesting apoptotic cells remained low over the course of the pulmonary immune response, and at all times exceeded the fraction of recruited monocytes ingesting apoptotic cells.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 References
 
Antibodies
The following monoclonal antibodies (mAbs) (clone; species isotype) were purchased from BD PharMingen (San Diego, CA): anti-CD11b (M1/70; rat IgG2b, {kappa}); anti-CD11c (HL3; Armenian hamster IgG1, {lambda}); anti-CD16/32 (2.4G2; rat IgG2b, {kappa}); anti-CD19 (1D3; rat IgG2a, {kappa}); anti-CD51 (RMV-7; rat IgG1); anti-CD90.2 (30-H12; rat IgG2b, {kappa}); anti-H-2Db (KH95; mouse IgG2b, {kappa}). Avidin-FITC and strepavidin-PerCP for use with biotinylated mAbs and the following controls (clone) were also purchased from BD PharMingen: control rat IgG2a, {kappa} (R35–95); control rat IgG2b, {kappa} (A95–1); control rat IgM (G155–228); anti-rat IgG2b (G15–337; mouse IgG2b, {kappa}); control hamster IgG1, {kappa} (A19–3); control hamster IgG3, {lambda} (A19–4). mAb specific for the Mø isoform of Ly-6c (clone ER-MP20; rat IgG2a, {kappa}) was obtained from Accurate Chemical and Scientific Corp (Westbury, NY). Anti-F4/80 (clone CI:A3–1; rat IgG2b) was purchased from Caltag Laboratories (Burlingame, CA). mAb against PSR (clone 217; mouse IgM) was a generous gift of Drs. Valerie Fadok and Peter Henson (National Jewish Medical Center; Denver, CO). Goat anti-mouse IgM was obtained from Jackson ImmunoResearch Laboratories (West Grove, PA). Anti-murine MerTK monoclonal (clone 108921, rat IgG1) and polyclonal (affinity-purified goat IgG) antibodies were obtained from R&D Systems (Minneapolis, MN). Normal goat IgG and rabbit anti-goat IgG (used as secondary antibody in MerTK staining) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA).

Mice
Pathogen-free inbred female mice were used at 8–14 wk of age in all experiments. C57BL/6 (H-2b) and BALB/c mice (H-2d) were purchased from Charles River Laboratories (Wilmington, MA) and Jackson Laboratory (Bar Harbor, ME), respectively. Mice were housed in the Animal Care Facility at the Ann Arbor Veterans Affairs Medical Center, which is fully accredited by the American Association for Accreditation of Laboratory Animal Care. Mice were fed standard animal chow (rodent lab chow 5001; Purina, St. Louis, MO) and chlorinated tap water ad libitum. This study followed a protocol approved by the Animal Care Committee of the local Institutional Review Board, and complied with the latest version of the Guide for the Care and Use of Laboratory Animals (National Academy of Sciences; www.nap.edu/readingroom/books/labrats/).

Experimental Design
Resident AMø were labeled in situ by intravenous administration of the long-lasting orange fluorescent vital dye PKH26 PCL, as previously described (14). Briefly, C57BL/6 mice that had been antigen-primed by IP injection of 1 x 108 SRBC in 0.5 ml normal saline 2–3 wk previously received 100 µl PKH26 PCL (Sigma, St. Louis, MO) (stock 1 x 10–3 M in ethanol, diluted 1:5 in Diluent B) by tail vein injection. This procedure labels mature phagocytic Mø throughout the body, with special efficiency in the lung, which receives the initial bolus of dye. Conversely, peripheral blood monocytes and their bone marrow precursors are not labeled, permitting their subsequent identification after recruitment to tissues (14–16).

In all experiments, alveolar mononuclear phagocytes were identified by light scatter characteristics using flow cytometry, distinguishing resident AMø as PKH26-positive (PKHpos) cells, and recruited monocytes as PKH26-negative (PKHneg) cells with the same light scatter characteristics. Three types of experiments were performed. First, phagocytosis of fluorescently-labeled apoptotic cells was measured in vitro; second, labeled apoptotic cells were instilled directly into the lungs, and phagocytosis was measured after recovery of Mø by BAL; and third, Mø receptor expression was analyzed by two- and three-color flow cytometry. In most experiments, BAL cells from individual mice were analyzed, as it was important to verify that each mouse received an adequate aliquot of PKH26 by tail-vein injection. In some experiments of receptor expression, the adequacy of PKH26 labeling of each mouse was first confirmed by flow cytometric analysis of PMø harvested from peritoneal lavage, and then BAL cells were pooled to obtain sufficient numbers of cells.

Induction of Pulmonary Immune Response
The particulate T cell–dependent antigen, sheep red blood cells (SRBC) (sheep 6115) (Colorado Serum, Denver, CO) were prepared and used as previously described (19). One day after intravenous injection of PKH26 PLC, SRBC-primed mice were antigen challenged by intratracheal injection of 5 x 10 8 SRBC in 50 µl normal saline. IT SRBC-challenge induces highly reproducible but self-limited lung inflammation characterized by vigorous mononuclear cell influx (19, 20). At various times after IT SRBC-challenge, mice were killed by asphyxia in a high CO2 environment, and bronchoalveolar lavage (BAL) and total cell counts were performed as previously described (19).

Thymocytes
Thymuses harvested from normal C57BL/6 mice were minced to yield a single cell suspension. Viable thymocytes were washed three times in Dulbecco's phosphate-buffered saline (PBS) (Invitrogen Life Sciences, Carlsbad, CA), suspended in PBS at a concentration of 2 x 107/ml, and incubated with the green fluorescent dye 5-chloromethylfluorescein diacetate (CMFDA) (Molecular Probes, Eugene, OR) according to the manufacturer's instructions. In some experiments, thymocytes from BALB/c mice (which express the MHC class I antigen H-2Dd) were used, so that target could be distinguished from phagocyte by staining for H-2Db expressed by C57BL/6 mice.

To induce apoptosis, CMFDA-labeled thymocytes were suspended at a concentration of 1–3 x 106/ml in RPMI 1640 medium containing 10% heat-inactivated FBS and a final concentration of 10–6 M dexamethasone (Sigma), and incubated for 6 h (7). This method results in a population of thymocytes in early apoptosis with little contamination by late apoptotic or necrotic cells (routinely 40–60 percent annexin-FITC positive, 12–18% annexin/PI double-positive, and < 2% trypan blue positive).

As a control target, CMFDA-labeled thymocytes were opsonized with 50 µl/ml of the rat monoclonal antibody CD90.2 for 30 min at 4°C, followed by mouse anti-rat mAb IgG for an additional 30 min. Opsonized thymocytes were washed with PBS and used immediately in the phagocytosis assay.

Phagocytosis Assays
Mø phagocytosis of apoptotic cells in vitro was analyzed as described by Jersmann and coworkers (21). BAL from individual mice were plated at 5 x 105 cells/well in sterile 48-well plates and allowed to adhere for 1–3 h at 37°C. Nonadherent cells were removed by gentle washing. Adherent Mø were incubated with 5 x 106 CMFDA-labeled apoptotic thymocytes or immunoglobulin-opsonized apoptotic thymocytes in 500 µl medium for 90 min (apoptotic) or 1 h (opsonized). Cells were extensively washed in ice-cold PBS to remove noningested thymocytes and were incubated in trypsin-EDTA at 4°C for 20 min and at 37°C for an additional 20 min. All aliquots of Mø released from the same plate were pooled and held on ice for analysis by flow cytometry immediately after the final wash. The adequacy of Mø release from the plate was confirmed in all experiments by direct inspection using an inverted microscope.

For in vivo experiments, 20 x 106 CMFDA-labeled apoptotic cells in 50 µl normal saline were administered intratracheally to SRBC-primed mice at various times after intratracheal SRBC challenge. Two hours later, mice were killed humanely by CO2 asphyxia and BAL was performed with 0.5 M EDTA in PBS. Cells were washed three times with 0.5 M EDTA in ice-cold PBS and immediately analyzed by flow cytometry.

Flow Cytometry
Cells freshly isolated by BAL were used to analyze expression of receptors potentially involved in clearance of apoptotic cells. Cells were washed twice in staining buffer (DIFCO, Detroit, MI), resuspended in 100 µl staining buffer, and incubated for 30 min at 4°C in the dark with labeled Abs diluted in 100 µl staining buffer. Final antibody concentrations were 1–2 µg/106 cells. Fc{gamma}R was blocked using mAb 2.4G2. After incubation, cells were washed twice in PBS, resuspended in 0.5 ml marking buffer, and analyzed immediately.

Flow cytometry was performed as previously described in detail (7) using a FACScan cytometer (Becton Dickinson, Mountain View, CA). Data were collected using CellQuest software (version 3.3; Becton Dickinson) on a PowerPC microcomputer (Apple, Cupertino, CA) and subsequently analyzed using Flo-Jo software (version 4.4.1; Tree Star, Ashland, OR). A minimum of 10,000 viable cells was analyzed to determine cell-surface receptor expression. To compare fluorescence intensities, we calculated the change in mean channel fluorescence ({Delta}MCF) between cells stained with specific antibody versus isotype control antibody.

Statistics
Data were expressed as mean ± SEM. Continuous ratio scale data were evaluated by unpaired Student's t test (for two samples) or ANOVA (for multiple comparisons) with post hoc analysis by the Tukey-Kramer test or by the two-tailed Dunnett test (for comparison to a single control condition). Statistical calculations were performed using Statview v 5.0 and Super ANOVA v 1.11 (SAS Institute, Inc., Cary, NC) on a Macintosh PowerPC G4 computer. Significant differences were defined as P < 0.05.


   RESULTS
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 References
 
Identification of Recruited Mononuclear Phagocytes in BAL
Resident AMø of normal mice were labeled brightly and uniformly by intravenously administered PKH26. The FL2 (orange) fluorescence intensity of AMø from mice receiving PKH26 did not overlap with that of resident AMø from normal untreated mice (Figure 1; compare Panel A with Panel B). Moreover, PKH26-labeled AMø of normal mice remained intensely stained for at least 13 d after a single intravenous administration, with virtually no loss of fluorescence intensity (data not shown). By contrast, prior treatment using PKH26 did not result in detectable staining of peripheral blood mononuclear cells during lung inflammation (Figures 1C and 1D). Flow cytometric analysis of BAL from normal mice that had been treated with PKH26 showed a distinct population of resident AMø, identified as cells of elevated side scatter (SSC) (Figure 1E). As anticipated, cells within this gate showed uniformly and high FL2 fluorescence, i.e., were PKH26-positive (PKHpos), but did not show high-level FL1 fluorescence (Figure 1F). This gate contained 92–97% of all BAL cells, the anticipated proportion of AMø based on differential cell counts. Conversely, PKHpos events were very uncommon outside these light scatter-defined gates (data not shown). Keeping instrument settings constant between experiments, we next applied identical light scatter gating to identify mononuclear phagocytes within BAL samples of intratracheally challenged mice (Figure 1G). At all times, this analysis disclosed a distinct population of PKH26-negative (PKHneg) cells that did not overlap in FL2 fluorescence with the PKHpos population (Figure 1H). Collectively, these data confirm that this technique permits clear distinction between resident AMø and recruited monocytes during immune lung inflammation, in agreement with previous studies using this technique (14–16).



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  		Figure 1. PKH26 treatment permits distinction between resident AMø and recruited monocytes during immune lung inflammation. (A and B) Baseline BAL data. AMø from unimmunized C57BL/6 mice were analyzed by flow cytometry using identical cytometer instrument settings, either without PKH treatment (A) or 24 h after injection of PKH26 by tail vein (B). Note the near absence of overlap in the FL2 fluorescence intensity (vertical axis) between the two populations. (C and D) Peripheral blood data. Mononuclear cells purified from peripheral blood of primed C57BL/6 mice 3 d after intratracheal SRBC challenge were analyzed by flow cytometry. Note minimal difference in FL2 fluorescence (vertical axis) between control mice not receiving PKH26 (C) and PKH26-treated primed mice (D). (E–H) BAL data during inflammation. SRBC-primed C57BL/6 mice received PKH26 by tail vein-injection, and 24 h later were either killed as a Day 0 control (E and F) or were challenged with 5 x 108 SRBC by the intratracheal route and assayed on various later days by flow cytometric analysis of BAL. Representative data are shown from a mouse 12 d after intratracheal challenge (G and H). (E and G) Light scatter data, as forward angle scatter (FSC) on the horizontal axis and side scatter (SSC) on the vertical axis. (F and H) Fluorescence data, as FL1 on the horizontal axis and FL2 (PKH26 staining) on the vertical axis. Note the appearance of PKHneg cells in the lower left-hand quadrant following immunization (H), and the absence of FL1+ cells (F and H).

 
Antigen challenge induced a marked increase in absolute numbers of PKHneg monocytes within BAL, from fewer than 100 cells/mouse at Day 0 to a peak at Day 3 of 0.5 ± 0.2 x 106 cells/mouse (mean ± SEM, n = 9) (Figure 2). This marked increase on Day 3 (> 8,000-fold above baseline) coincided with the peak of alveolar inflammation and of total Mø numbers previously described in this model system (19). Thereafter, numbers of PKHneg monocytes decreased, but remained slightly above baseline through 12 d after intratracheal challenge. By contrast, absolute numbers of PKHpos resident AMø in BAL did not change significantly throughout the first 6 d of the response (Figure 2), although there was a nonsignificant trend toward higher numbers at Days 9 and 12 (P > 0.5 at all time points compared with Day 0, ANOVA with Dunnett post hoc testing).



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  		Figure 2. Kinetics of mononuclear cell recruitment during immune lung inflammation. SRBC-primed mice received PKH26 by tail vein-injection and were challenged by the intratracheal route 24 h later with 5 x 108 SRBC. At the indicated days after intratracheal SRBC-challenge, mice underwent BAL, and absolute cell counts of PKHpos Mø (black squares) and PKHneg Mø (open circles) cells were calculated. Results are mean ± SEM of 3–10 mice per time point assayed individually in at least three separate experiments per time-point.

 
To characterize the two Mø populations further, BAL cells from primed mice were stained on various days after intratracheal challenge with mAbs against a variety of leukocyte antigens and analyzed by flow cytometry (Figure 3). PKHpos Mø were only weakly positive for F4/80 (Figure 3A), in agreement with previous data on resident murine AMø (22), whereas PKHneg Mø strongly expressed F4/80 (Figure 3B) (% fluorescence-positive: 2.5 ± 0.9% for PKHpos Mø versus 28.2 ± 2.6% for PKHneg Mø; {Delta}MCF: 41.6 ± 3.7 for PKHpos Mø versus 121.1 ± 13.4 for PKHneg Mø; mean ± SEM of eight mice assayed individually on Day 3 after intratracheal challenge; P < 0.05, unpaired t test). PKHpos Mø stained weakly with mAb ER-MP20 (Figure 3C), whereas PKHneg Mø were stained strongly (Figure 3D), as would be anticipated for relatively immature mononuclear phagocytes (23) (% fluorescence-positive: 1.1 ± 0.5 for PKHpos Mø versus 77.2 ± 3.4 for PKHneg Mø; {Delta}MCF: 52.0 ± 6.1 PKHpos Mø versus 328.3 ± 15.2 PKHneg Mø; mean ± SEM of eight mice assayed individually on Day 3 after intratracheal challenge; P < 0.05, unpaired t test). Negligible staining was seen within either the PKHpos or PKHneg Mø populations for the lymphocyte antigens TCR-ß (Figures 3E and 3F) and CD19 (Figures 3G and 3H), which are pathognomonic of T cells and B-cells, respectively. As anticipated from our previous data on resident murine AMø (7), PKHpos Mø were brightly positive for CD11c and showed minimal staining for CD11b, even late after intratracheal challenge (Figure 3G), whereas PKHneg Mø remained mostly positive for CD11b alone (Figure 3H). Almost no cells within the Mø gate stained brightly for the granulocyte marker Gr-1 at any time (data not shown). To determine whether our gating strategy captured all BAL mononuclear phagocytes, we also examined cells of lower orthogonal light scatter ("lymphocyte gate"). Staining for F4/80 and ER-MP20 was essentially absent from this gate, whereas the majority of cells did stain for one or more typical lymphocyte markers such as CD3, CD19 and TCR-ß (data not shown). These data collectively indicate that light scatter-gating accurately identifies a population of alveolar cells highly enriched for mononuclear phagocytes, with minimal contamination by other cell types, and that the population we analyzed contains the vast majority of alveolar mononuclear phagocytes.



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  		Figure 3. Characterization of mononuclear phagocyte subpopulations during immune lung inflammation. SRBC-primed mice received PKH26 by tail vein-injection and were challenged by the intratracheal route 24 h later with 5 x 108 SRBC. At various later times, alveolar cells were harvested by BAL and cells within light scatter-defined gates were analyzed for the indicated antigens by flow cytometry. (A, C, E, G, and I) PKHpos Mø. (B, D, F, H, and J) PKHneg Mø. (A–H) Representative histograms from individual mice, 3 d after intratracheal challenge. Horizontal axis, log FL1 fluorescence; dotted line, isotype control; solid line, specific staining. (I and J) Representative two-color fluorescence plots from a single mouse, 12 d after intratracheal challenge. Horizontal axis, log FL1 fluorescence (CD11b); vertical axis, log FL3 fluorescence (CD11c). Quadrants indicate specific staining for each cell type.

 
In Vitro Phagocytosis by Alveolar Mononuclear Cells
We next investigated the capacity of the two Mø populations to ingest fluorescently-labeled apoptotic cells in vitro. BAL was harvested from normal mice or from intratracheally challenged mice to permit isolation of Mø by adherence. In control experiments, we verified that this initial adherence step did not significantly change the ratio of PKHneg Mø to PKHpos (12.2 ± 4.1% PKHneg Mø pre-wash versus 8.8 ± 1.5%. PKHneg Mø post-wash; P = 0.46, Student's t test; mean ± SEM of triplicate samples in each of five individual experiments). Additional control experiments confirmed that in vivo labeling of AMø using intravenously administered PKH26 PCL had a minimal effect on phagocytic ability (PKH26 treatment 94.7 ± 7.8% of control; P = 0.76, Student's t test; mean ± SEM in three individual experiments).

Mø were next co-cultured with CMFDA-labeled apoptotic thymocytes. Uningested thymocytes were removed by standardized washing, and then Mø were released from the dishes using cold EDTA and analyzed by microscopy and two-color flow cytometry. The adequacy of Mø recovery from the dishes was confirmed in all experiments by direct inspection using an inverted microscope, with additional washes if necessary.

Using light and fluorescent microscopy, we first confirmed that fluorescent targets were inside Mø, and not merely adherent (Figures 4A and 4B). Flow cytometry showed that within both PKHpos and PKHneg populations, Mø that had ingested labeled thymocytes were easily distinguishable due to their very increased FL1 fluorescence (Figures 4C and 4D). Moreover, the percentage of FL1-positive Mø in the PKHpos population in normal mice as determined by this method agreed with our previous data on resident murine AMø using direct inspection of microscopy slides (7). As a more definitive test that our assay measured only intracellular thymocytes, we also performed a control phagocytosis experiment to quantify bound but uningested thymocytes, based on differences in MHC class I antigen expression between Mø and thymocytes. We used BAL cells from C57/BL/6 mice (H-2Db) and apoptotic thymocytes from BALB/c mice (H-2Dd), and stained the Mø (without permeabilization) after the final wash step using biotinylated anti-H-2Dd plus streptavidin PerCP (an FL3 reagent). This reagent would stain any extracellular adherent thymocytes but not ingested thymocytes. Results confirmed minimal contribution of adherent uningested thymocytes (2.8 ± 1.1% of CMFDA-positive Mø were H-2Dd positive; mean ± SEM of four mice assayed individually in two separate experiments). Thus, this assay system appears to measure ingested apoptotic cells accurately.



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  		Figure 4. Phagocytosis of apoptotic thymocytes by recruited and resident mononuclear phagocytes in vitro. At various days after intratracheal SRBC challenge of primed PKH-labeled mice, cells were recovered by BAL, washed, and allowed to adhere to plastic plates. CMFDA-labeled apoptotic thymocytes (A–E) or CMFDA-labeled apoptotic thymocytes opsonized by IgG (F) were added to the adherent Mø at a ratio of 10:1 and incubated for 90 min. Slides were vigorously washed to remove external thymocytes and Mø were released into suspension using trypsin/EDTA, and analyzed by fluorescence microscopy (A and B) or flow cytometry (C–F). (A and B) Photomicrographs of Mø containing internalized apoptotic thymocytes; epifluorescence illumination (A); phase contrast (B). Note the distinction between the uningested green thymocyte not associated with Mø (white arrow), and a PKHpos Mø containing two ingested yellow thymocytes (yellow arrowheads). (C and D) Representative dot plots from individual mice incubated either without (C) or with (D) CMFDA+ apoptotic thymocytes. Note appearance of FL1+ cells in both right-hand quadrants of D, indicating Mø that have ingested apoptotic thymocytes. (E) Kinetics of Mø phagocytosis of apoptotic thymocytes in vitro. Black squares, PKHpos Mø; open circles, PKHneg Mø. (F) Kinetics of phagocytosis of opsonized apoptotic thymocytes in vitro. Dark bars, PKHpos Mø; light bars, PKHneg Mø; ND = not done. Results in E and F are mean ± SEM of 3–9 mice assayed individually in at least three independent experiments; *P < 0.05, unpaired t test compared with PKHneg Mø at the same time-point.

 
Kinetic analysis over the pulmonary immune response disclosed a marked difference in the efficiency of apoptotic cell phagocytosis between the two mononuclear phagocyte subpopulations. At all time points after intratracheal challenge, the percentage of phagocytic cells among PKHpos Mø was greater than among PKHneg Mø (Figure 4E). The fraction of phagocytic PKHpos Mø remained statistically unchanged during the immune response, ranging between 27.6 and 44.9% (P > 0.5 at all time points compared with Day 0, ANOVA with Dunnett post hoc testing). Phagocytosis by PKHneg Mø could not be measured at Day 0, due to the extreme paucity of that cell population, but thereafter ranged between 13 and 21%, also not changing statistically (P > 0.5, ANOVA with Tukey-Kramer post hoc testing). However, both PKHpos and PKHneg populations avidly ingested immunoglobulin-opsonized apoptotic cells (Figure 4F), confirming their capacity to ingest targets of identical size using an alternative recognition pathway (Fc{gamma}R-mediated). These data demonstrate the ability of our assay system to detect high degrees of phagocytosis when present. The observed avid Fc{gamma}R-mediated phagocytosis also argues strongly against the possibility that lung Mø were already stuffed with apoptotic cells they had ingested previously in vivo, an explanation we believed highly unlikely based on their morphologic appearance.

In Vivo Phagocytosis
We next examined Mø clearance of apoptotic cells from the alveolar spaces. For this purpose, we administered CMFDA-labeled apoptotic thymocytes by the intratracheal route to SRBC-primed, PKH-treated mice that had been challenged at various earlier times with intratracheal SRBC, and 2 h later harvested BAL and analyzed Mø by flow cytometry. Phagocytosis of apoptotic thymocytes was seen in both Mø populations, and the percentage of phagocytic (FL1-positive) Mø in both populations agreed with results of the in vitro phagocytosis assay. PKHneg Mø again ingested fewer than did PKHpos Mø at all time points, although the difference between the two cell types was not significant (Figure 5) (P > 0.05, unpaired t test). When analyzed over time after intratracheal SRBC challenge, the percentage of phagocytic Mø did not change significantly in PKHpos Mø (P > 0.5 at both time points compared with Day 0, ANOVA with Dunnett post hoc testing), or in PKHneg Mø (P > 0.5, Day 5 versus Day 12, unpaired t test).



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  		Figure 5. Kinetics of phagocytosis of apoptotic thymocytes in vivo. SRBC-primed mice received PKH26 by tail vein-injection and were challenged by the intratracheal route 24 h later with 5 x 108 SRBC to induce lung inflammation. At the indicated days after intratracheal challenge, mice each received 20 x 106 CMFDA-labeled apoptotic thymocytes by the intratracheal route, and were allowed to recover from anesthesia. Two hours later, mice were killed, alveolar cells were recovered by BAL, and the percentage of phagocytic Mø in the PKHpos and PKHneg subsets was determined by two-color flow cytometry. Black bars, PKHpos Mø; white bars, PKHneg Mø. Results are mean ± SEM of 3–8 mice in at least two independent experiments per time-point. Day 0 indicates SRBC-primed C57BL/6 mice that had received PKH26 and labeled apoptotic thymocytes, but no intratracheal SRBC. n.d., not determined.

 
Because these in vivo experiments used large numbers of thymocytes per mouse, one potential confounding factor might be the presence of thymic Mø carried over in the thymocytes population. Such cells would have been CMFDA-labeled ex vivo before administration, and hence might be misidentified as phagocytosis-positive PKHneg Mø. To verify that our assay measured only phagocytosis by recruited PKHneg host monocytes, in a control experiment (Day 5 after intratracheal challenge) we again used thymocytes from mice that differed at the class I MHC locus. BAL Mø from C57BL/6 mice (which express H-2Db) were incubated with CMFDA-labeled apoptotic thymocytes from BALB/c mice (which express H-2Dd). We performed the same in vivo phagocytosis assay, except that cells were also stained with biotinylated anti-H-2Db and strepavidin-PerCP. Virtually no FL3-negative cells were identified within the Mø gates, and identical percentages of phagocytosis were seen whether or not analysis was restricted to FL3-positive cells. These data exclude carryover of thymic Mø as a source of error in this assay.

We also considered the possibility that the intratracheal instillation might be inhomogeneous in individual mice, and that some Mø might be negative because they were not exposed to apoptotic cells in vivo. To address this possibility, we performed an additional experiment in which we mixed unstained apoptotic thymocytes with zymosan-FITC, instilled the mixture by the intratracheal route, and counted only FITC-containing Mø, using our previously described microscopic assay of phagocytosis (7). A preliminary experiment had shown that resident AMø from normal mice were able to ingest apoptotic thymocytes and FITC-labeled zymosan simultaneously at the same rate as they ingested each particle independently (see Table E1 in the online supplement), and that virtually all AMø avidly ingested zymosan. Therefore, we anticipated that ingestion of zymosan-FITC could be used as a surrogate for exposure to the intratracheal bolus. Indeed, 100% of Mø containing apoptotic thymocytes also were FITC-positive, and 22.8 ± 4.6% of FITC-positive Mø contained apoptotic thymocytes (mean ± SEM, n = 3), in close agreement with our previous results; note that because this experiment was counted using microscopy, we did not attempt to distinguish PKH positivity. Thus, inhomogeneity of apoptotic cell deposition within the lungs does not appear to explain the relatively low fraction of alveolar mononuclear phagocytes ingesting apoptotic cells in vivo.

Surface Molecule Expression
Finally, we analyzed expression of three Mø surface receptors previously implicated in the phagocytosis of apoptotic cells. In hematopoietic cells, the {alpha}V integrin (CD51) forms a receptor for vitronectin together with either ß3 (CD61) or ß5 integrin chains. Through use of blocking mAb or inhibition with the tetrapeptide RGDS, experimental models utilizing human peripheral blood–derived monocytes or murine Mø cell lines have shown that binding of vitronectin to thrombospondin on the apoptotic cell surface is important for phagocytosis by these cell types (24, 25). Nevertheless, we found that PKHpos AMø did not express CD51 at any time during the pulmonary immune response (Figures 6A and 6C, left-hand panels) in agreement with our previous data on resident AMø from untreated mice (7). By contrast, PKHneg Mø strongly expressed CD51 on Day 3 after IT SRBC-challenge (Figure 6B, left-hand panel), with a mean intensity of fluorescence that fell progressively thereafter (Figure 6C, left-hand panel). However, as we have previously shown for normal resident AMø and PMø from untreated mice (7), in separate functional experiments, the integrin-blocking tetrapeptide RGDS did not decrease phagocytosis by either PKHpos Mø or PKHneg Mø (data not shown). These results imply that clearance of apoptotic lung lymphocytes by murine lung Mø during immune inflammation does not depend on vitronectin binding.



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  		Figure 6. Expression of receptors important for apoptotic cell clearance. SRBC-primed mice received PKH26 by tail vein-injection, were challenged by the intratracheal route 24 h later with 5 x 108 SRBC, and were assayed on various later days for expression of CD51 (left-hand column), PSR (middle column), and MerTK (right-hand column) by flow cytometric analysis of BAL. (A and B) Representative flow cytometry data from individual mice 6 d after intratracheal challenge, gated on (A) PKHpos Mø and (B) PKHneg Mø. (C) Kinetics of receptor expression. Black bars, PKHpos Mø; white bars, PKHneg Mø. Results are expressed as change in mean fluorescence channel number ({Delta} MCF) from isotype control, as mean ± SEM of 3–5 mice per time-point, assayed individually. *Significantly different from other Mø population, P < 0.05, unpaired t test. Note difference in scale of {Delta} MCF between different receptors.

 
The stereospecific PSR appears to be central to apoptotic cell recognition (12). We found that PKHpos Mø and PKHneg Mø had similar low PSR expression that changed little during the immune response (Figure 6, center panels).

PKHpos Mø abundantly expressed the receptor tyrosine kinase MerTK (Figures 6A and 6C, right-hand panels), which we have recently shown is involved in apoptotic cell phagocytosis by resident murine AMø (10). The intensity of MerTK expression by PKHpos Mø increased markedly during resolution of lung inflammation, as demonstrated by {Delta}MCF (Figure 6C, right-hand panel). Importantly, PKHneg Mø in the same samples barely expressed MerTK at any time-point (Figures 6B and 6C, right-hand panels). Antibody against MerTK significantly inhibited in vitro phagocytosis of apoptotic cells by PKHpos Mø (anti-MerTK 21.4 ± 2.6% phagocytic Mø versus isotype control 45.3 ± 2.7% phagocytic Mø, n = 9, P < 0.0001), but had no significant effect on PKHneg Mø (anti-MerTK 20.8 ± 4.2% phagocytic Mø versus isotype control 20.6 ± 3.1% phagocytic Mø, P = 0.81).


   DISCUSSION
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
References
 
The results of this study define the clearance of apoptotic lymphocytes by murine alveolar mononuclear phagocytes during antigen-induced lung inflammation. The principal conclusion is that among both resident AMø and recruited monocytes, the percentage of individual alveolar mononuclear phagocytes ingesting apoptotic lymphocytes remains low at all times relative to ingestion of immunoglobulin-opsonized targets. This conclusion is based on the following findings: (1) the fraction of PKHpos AMø ingesting apoptotic cells did not increase significantly from low baseline levels, in vitro or in vivo; and (2) despite a rapid influx of PKHneg recruited monocytes that greatly yet transiently increased total numbers of mononuclear cells in the alveoli, recruited monocytes displayed an even lower rate of apoptotic cell ingestion in vitro than PKHpos resident AMø at all times. The relative deficiency in apoptotic cell ingestion by recruited monocytes was specific, as shown by the avid ingestion of immunoglobulin-opsonized particles, and was associated with low expression of the receptor tyrosine kinase MerTK. Collectively, these data explain our previous observation that apoptotic lymphocytes, which are easily detected in the alveoli in this experimental model system (18), are present due to inefficient clearance by phagocytes in the alveolar space. We also found that even though the vitronectin receptor chain CD51 was abundantly expressed by recruited monocytes (but not by resident AMø throughout the immune response), phagocytosis of apoptotic cells by both populations of alveolar mononuclear phagocytes was not inhibited by the integrin-blocking tetrapeptide RGDS. These novel findings have important implications for the clearance of CD8+ T cells following viral pneumonias (26), and motivate additional investigation of how lung mononuclear cells contribute to resolution of inflammation.

Our results showing the accurate distinction of recruited monocytes from resident AMø using PKH26 confirm and extend those of Maus and colleagues, who pioneered this technique to label lung Mø in vivo (14–16). Monocytes have been studied to a much lesser degree in murine models than in humans, due in part to the marked difficulty in obtaining pure populations from the peripheral blood of animals as small as mice. Moreover, there have been no clear-cut alternative methods to distinguish unambiguously between recently recruited monocytes and resident AMø Mø, either morphologically or by surface markers (4). By permitting the unique contribution of recruited monocytes to immune and inflammatory responses to be defined, this technique is an important innovation that could be applied to other organs. We believe that analysis by flow cytometry is essential to provide reproducible results; preliminary experiments using immunofluorescence microscopy convinced us that the human eye is unable to distinguish reliably between the fluorescence intensity of PKHpos Mø and PKHneg Mø (unpublished observation). The lung appears to be a particularly favorable tissue for use of this technique. We noted that staining of AMø was consistently of greater intensity than that of PMø in the same mice (unpublished observation), likely due to the high first-pass effect of an intravenous injection on the pulmonary vasculature.

An important facet of our data is the insights they give into the differing fates of resident AMø and recruited monocytes during immune lung inflammation. Particle-laden mature AMø have been shown previously to migrate to regional lymph nodes (27), but the degree to which this migration depletes AMø numbers within the lung has not been investigated. The finding that overall numbers of resident AMø did not change significantly over the course of this immune response implies that the number of AMø leaving the lungs during the pulmonary immune response is relatively small. Support for this conclusion comes from preliminary experiments which show very few PKHpos Mø in mediastinal lymph nodes in this model system (unpublished observation). Emigration of resident AMø may also have been balanced by transmigration into the alveoli of interstitial lung Mø, which should also have been stained by PKH26. This possibility is supported by the trend toward increased numbers of PKHpos Mø seen at our later two time-points. Conversely, our kinetic data indicate that the vast majority of monocytes recruited to the lungs during pulmonary immune responses do not become long-term residents of the alveolar spaces. Although we did not directly investigate the mechanism for the rapid decrease in numbers of monocytes after Day 3, it is likely to result primarily from emigration (Ref. 28, and unpublished data). Moreover, PKHneg Mø remained relatively small in size, as shown by low forward light scatter, and continued to express surface antigens such as F4/80 and ER-MP20 (data not shown), consistent with an immature phenotype. Additional studies will be needed to explicate fully Mø population dynamics during lung inflammation.

These results also provide a novel in vivo perspective on the important literature investigating phagocytic capacity of human monocyte-derived Mø during in vitro maturation. We showed that at least some murine monocytes develop phagocytic capacity by Day 3 after intratracheal challenge, which we believe from more detailed kinetic analyses of leukocyte trafficking in this model system (Refs. 17 and 19, and unpublished observations) indicates ~ 24–48 h of residence in the lungs. By contrast, freshly isolated human monocytes did not ingest apoptotic cells at all, and can require up to a week in culture to gain that function (13). For ethical reasons, it would be difficult to use this staining technique in humans to determine whether these disparate results reflect species differences or simply more rapid development of at least some phagocytic capacity in vivo; we favor the latter interpretation. Our finding that recruited murine monocytes did not further upregulate their phagocytic capacity over 12 d of lung inflammation could be interpreted to indicate that that individual monocytes may have stayed in the lungs for only a brief period. Our results in mice also differ somewhat from those of Newman and colleagues, who showed a more pronounced increase from baseline phagocytosis of neutrophils by alveolar mononuclear phagocytes of rabbits treated with intratracheal acid (6). This apparent disparity could be due to a greater degree of tissue injury in their model system or the difference in apoptotic target, as Mø have unique mechanisms restricted to recognition of apoptotic neutrophils (29).

The finding that recruited monocytes express extremely low levels of the Tyro3 receptor kinase family member MerTK provides one potential mechanism for their low rate of apoptotic cell phagocytosis. Several pieces of evidence indicate a central role for MerTK in apoptotic cell clearance (30). Transgenic mice exclusively expressing a kinase-deficient form of Mer (Merkd), which interferes with MerTK surface expression, clear apoptotic cells poorly and develop auto-immunity as they age (2, 31, 32). Genetic deficiency or mutation of MerTK leads to retinitis pigmentosa in rats and humans (30) due to defective clearance of apoptotic bodies by retinal pigment epithelial cells. We have shown that polyclonal antibody against MerTK blocks phagocytosis (but not adhesion) of apoptotic cells by murine Mø, including AMø (10); as anticipated, the current data extend that finding to mAb against MerTK and this flow cytometry-based assay. The other members of this receptor kinase family, Axl and Tyro3, have also been shown to be involved in apoptotic cell clearance in vivo (30, 33), but little is known about their contribution to ingestion by resident Mø from specific tissues. Whether murine monocytes also lack expression of other Tyro3 family members is under current investigation.

We carefully considered a variety of technical limitations that could detract from this analysis. First, it is unlikely that our assay conditions missed very rapid uptake and ingestion of apoptotic cells, as we have previously performed extensive kinetic analysis (15 min to 8 h of co-incubation) of Mø adhesion and phagocytosis using a microscopic assay system (7, 10). Moreover, CMFDA should be retained within Mø even if the ingested cells were digested rapidly. Second, we think it highly unlikely that some unique feature of SRBC challenge specifically reduces Mø ingestion of apoptotic cells, as our current data during inflammation agree closely with our current and previous results in normal mice. Third, because we have shown that a substantial fraction of murine alveolar lymphocytes are apoptotic throughout the course of this response (18), we cannot exclude the possibility that the relatively low capacity of resident PKHpos AMø to ingest apoptotic cells reflects the effect of chronic low level ingestion. Previous ingestion of apoptotic neutrophils has been shown to decrease the subsequent capacity of rat bone marrow-derived Mø to ingest apoptotic neutrophils in vitro (34). However, this critique would obviously not apply to PKHneg monocytes. Overall, we are confident that our results accurately mimic the behavior of murine lung mononuclear phagocytes in both the normal state and in a variety of infectious and inflammatory diseases.

If both resident AMø and especially recruited monocytes have limited capacity to ingest apoptotic leukocytes, how then are they cleared during resolving inflammation? The simple answer in this model system (18) is that they are cleared slowly and inefficiently, which is why they are easily detected. We suspect that similar delayed clearance occurs following viral pneumonias (26). It is also possible that apoptotic leukocytes are cleared in part by pulmonary airway epithelial cells, which can ingest apoptotic eosinophils in vitro (35). Epithelial cells appear to have an even lower efficiency per individual cell than do AMø, but if type II alveolar epithelial cells show the same phagocytic behavior as the A549 cell line (36), their massive total numbers argue that they might make a sizeable net contribution. We did not specifically measure recovery of uningested thymocytes in our in vivo experiments, but it was clear from hemocytometer counts and inspection of the ungated flow cytometric results that the vast majority of apoptotic cells were not ingested during that short assay.

In summary, we have shown that both resident AMø and, to a lesser degree, recruited monocytes, can ingest apoptotic lymphocytes during antigen-induced immune lung inflammation, but that the phagocytic fraction of both cell types is low, and does not increase markedly over time. When the overall phagocytic capacity of resident AMø versus recruited monocytes for apoptotic cells is estimated as the product of the observed absolute numbers per mouse and the percent phagocytic cells of each subset, it becomes clear that resident AMø contribute the overwhelming amount of ingestion at all times except Day 3. These results emphasize the continued need to study how the interaction of apoptotic cells with AMø modulates both ongoing inflammation and host defense of the lungs.


   Acknowledgments
 
The authors thank Drs. Geoffrey J. Bellingan, Ian Dransfield, Valerie Fadok, Peter Henson, and William Vandivier, and all the members of the Ann Arbor VA REAP for helpful suggestions and discussion; Joyce O'Brien for secretarial support; and Dr. Antonello Punturieri for critiquing the manuscript.


   Footnotes
 
Portions of this work have been presented previously at the International Conference of the American Thoracic Society, Seattle, WA, May 19, 2003, and have been published in abstract form (Am J Respir Crit Care Med 2003; 167:A491).

This study was supported by RO1 HL56309 and R37 HL34788 from the USPHS; by Merit Review funding, a Career Development award, and a Research Enhancement Award Program (REAP) grant from the Department of Veterans Affairs; and by funding from the Michigan Life Sciences Initiative.

This article has an online supplement, which is accessible from this issue's table of contents at www.atsjournals.org

Conflict of Interest Statement: J.H.J. has no declared conflicts of interest; D.J.L. has no declared conflicts of interest; B.H. has no declared conflicts of interest; J.S. has no declared conflicts of interest; and J.L.C. has no declared conflicts of interest.

Received in original form March 31, 2004

Received in final form November 7, 2004


   References
Top
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 Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 

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