American Journal of Respiratory Cell and Molecular Biology

In the acute respiratory distress syndrome, recruitment of peripheral blood monocytes results in expansion of the total pool of resident alveolar macrophages. The fate of resident macrophages, or whether recruited monocytes are selectively eliminated from the alveolar airspace or differentiate into resident alveolar macrophages during the resolving phase of inflammation, has not been determined. Here, we analyzed the kinetics of resident and recruited macrophage turnover within the alveolar airspace of untreated and LPS-challenged mice. Using bone marrow chimeric CD45.2 mice that were generated by lethal irradiation of CD45.2 alloantigen-expressing recipient mice and bone marrow transplantation from CD45.1 alloantigen-expressing donor mice, we employed a flow cytometric approach to distinguish recipient from donor-type macrophages in bronchoalveolar lavage fluids. Our data show that resident alveolar macrophages of untreated chimeric CD45.2 mice are very slowly replaced by constitutively immigrating CD45.1 positive monocytes, resulting in a replacement rate of ∼ 40% by 1 yr. In contrast, more than 85% of the resident CD45.2 positive alveolar and lung homogenate macrophages were exchanged by donor CD45.1-expressing macrophages within 2 mo after treatment with Escherichia coli endotoxin (LPS). Importantly, fluorescence-activated cell sorter analysis of increased annexin V binding to both recipient and donor-type macrophages revealed increased apoptotic events to underlie this endotoxin-driven inflammatory macrophage turnover. Collectively, the data show that under baseline conditions the alveolar macrophage turnover exhibits very slow kinetics, whereas acute lung inflammation in response to treatment with LPS triggers a brisk acceleration of recruitment of monocytes that replace the resident alveolar macrophage population.

The acute respiratory distress syndrome (ARDS) is characterized by a rapid increase of alveolar neutrophils with a relatively delayed increase in the population of alveolar macrophages. Acute lung injury developing in patients with ARDS correlates strongly with increased concentration of CCL2 in bronchoalveolar lavage (BAL) fluid (BALF). CCL2 is a major monocyte chemoattractant centrally involved in the recruitment of peripheral blood (PB) monocytes that is associated with acute lung inflammation (13). Recent evidence in a mouse model of acute lung inflammation indicates that a genetic deletion or treatment with neutralizing antibodies for CCR2, the primary receptor for CCL2 expressed on circulating monocytes, abrogates both recruitment of PB monocytes and prevents neutrophilic lung inflammation in mice (4). These data suggest the hypothesis that inflammatory monocyte trafficking to the lung, which is regulated by the CCL2/CCR2 axis, is a critical event in lung inflammation and injury that is associated with gram-negative bacterial endotoxemia.

Most recently, we found that monocytes that are recruited to the alveolar space under acute inflammatory conditions displayed a prominent proinflammatory genotype, as manifested by increased steady-state mRNA levels of the proinflammatory cytokine TNF-α; the neutrophil chemoattractants macrophage inflammatory protein (MIP)-2, KC, and IP-10; the pattern recognition receptors and signaling molecules TLR4 and MD1; and various lysosomal cysteine proteases involved in extracellular matrix degradation and tissue remodeling processes (5). Inflammatory monocyte recruitment to the lung may therefore amplify the overall lung inflammatory response to bacterial pathogen-associated molecular patterns (PAMP) by both releasing proinflammatory mediators within the alveolar micromilieu and by releasing ECM-degrading proteolytic activities, which may potentiate acute lung injury. Interestingly, both CCR2 deletion and neutralization of CCL2 attenuate TNF-α and CINC levels within the alveolar airspace of mice and rats (6, 7). On the other hand, blockade of inflammatory monocyte trafficking in a mouse model of pneumolysin-induced acute lung injury strongly delayed the repopulation of the pneumolysin-depleted alveolar macrophage pool, with possible side-effects on the lung defense against pneumococcal colonization (8). Taken together, a balanced manipulation of monocyte trafficking to the acutely inflamed lung may be beneficial in modulating the inflammatory response to bacterial toxins. However, such maneuvers require a precise knowledge about the interplay between newly alveolar recruited monocytes and resident alveolar macrophages and the fate of either population during resolving inflammation, since they may disrupt innate host defense mechanisms.

In the current study, we determined the turnover of the resident alveolar macrophage pool under baseline conditions and in response to Escherichia coli LPS challenge. To facilitate the clear-cut discrimination between resident and newly recruited monocytes/macrophages under baseline versus inflammatory conditions, we generated chimeric mice by transplanting donor CD45.1 bone marrow cells into lethally irradiated CD45.2 recipient mice. Flow cytometric assessment of baseline alveolar macrophage exchange kinetics in these chimeric CD45.2 mice detected a very slow turnover of the resident macrophage population under noninflammatory conditions. In contrast, acute lung inflammation triggered a strongly accelerated, increased and persistent exchange of the recipient-type resident alveolar macrophage pool against donor-type macrophages.

Animals

Recipient C57BL/6J mice expressing the CD45.2 alloantigen (Ly5.2 protein tyrosine phosphatase, PTP) (9) on the cell surface of circulating leukocytes were obtained from Charles River (Sulzfeld, Germany). Donor B6.SJL-Ptprca mice (C57BL/6) expressing the CD45.1 alloantigen (Ly5.1 PTP) (9) on circulating leukocytes were purchased from Jackson Laboratories (Bar Harbor, ME). Animals were housed under conventional conditions and were used for the described experiments at the age of 8–10 wk and between 18–21 g body weight. This animal study was approved by the local government committee.

Reagents

E. coli endotoxin (E. coli LPS, serotype O111:B4) was purchased from Sigma (Deisenhofen, Germany). Flow cytometric detection of the CD45.1 alloantigen (Ly5.1) expression on donor-type PB leukocytes and BAL fluid mononuclear phagocytes was performed with either fluorescein isothiocyanate (FITC)- or phycoerythrine (PE)-labeled mouse anti-mouse CD45.1 monoclonal antibody (clone A20, isotype mouse IgG2a). CD45.2 alloantigen (Ly5.2) expression on recipient-type PB leukocytes and BAL fluid mononuclear phagocytes was detected using FITC-labeled mouse anti-mouse CD45.2 monoclonal antibody (clone 104, isotype mouse IgG2a). All antibodies and isotype-matched irrelevant control antibodies were obtained from BD Biosciences (Heidelberg, Germany). Phycoerythrin-labeled annexin V was also purchased from BD Biosciences.

Isolation of CD45.1-Positive Bone Marrow Cells and Generation of Chimeric CD45.2 Alloantigen-Expressing Recipient Mice

Bone marrow cells were isolated under sterile conditions from the tibias and femurs of sex-matched CD45.1 donor mice, following recently published protocols (6, 10). Briefly, tibias and femurs were flushed with sterile RPMI/10% FCS and a single cell suspension was prepared from the bone marrow isolates, filtered through 70-μm and 40-μm nylon meshes (BD Biosciences) to remove cell aggregates, and bone marrow cell suspensions were then washed in Leibovitz L15 medium (Gibco, Karlsruhe, Germany) before transplantation. Recipient CD45.2 alloantigen-expressing C57BL/6 mice received 12 Gy of a total body irradiation using a 60Co source split into two doses (8 Gy and 4 Gy) separated by a 3-h interval to reduce gastrointestinal toxicity (10). A total of 1 × 107 CD45.1 donor bone marrow cells suspended in Leibovitz L15 medium were transplanted via lateral tail vein injections into sedated CD45.2 recipient mice. Resulting chimeric CD45.2 recipient mice characterized by a CD45.1-positive hematopoietic system and a CD45.2-expressing resident alveolar macrophage pool were then housed under specific pathogen–free (SPF) conditions with free access to autoclaved food and water.

Isolation of PB Leukocytes and Alveolar Macrophages, and Preparation of Lung Homogenate Macrophages

Mice were killed with an overdose of isoflurane (Forene; Abbott, Wiesbaden, Germany). PB was collected from the inferior vena cava, and BAL was performed as previously described (3, 10, 11). Briefly, after mice were killed the trachea was exposed by a midline incision. A shortened 21-G cannula was inserted into the trachea and fixed tightly. BAL was done by initially instilling a 300-μl aliquot of 4°C cold PBS solution (supplemented with 2 mM EDTA; Versen, Berlin, Germany) into the mouse lungs followed by gentle aspiration, followed by instillation of a 400-μl aliquot of cold PBS and aspiration, and then instillations of 500-μl aliquots each of cold PBS into the lungs, until a total BALF volume of 5 ml was collected and centrifuged for 10 min at 1,200 rpm at 4°C (Heraeus, Hanau, Germany). The cell pellet was resuspended in RPMI/10%FCS, and total cell numbers of alveolar macrophages in BAL fluids of untreated or LPS-challenged chimeric CD45.2 mice were quantified using overall morphologic criteria including cell size and shape of nuclei on Pappenheim-stained cytocentrifuge preparations.

In selected experiments, the LPS-induced macrophage turnover was also analyzed in lung homogenates of LPS-treated chimeric CD45.2 mice. Briefly, mice were killed with an overdose of isoflurane and the lungs were subjected to BAL as described above. The thorax was carefully opened and the lungs were perfused via the right ventricle with Hanks' balanced salt solution (HBSS) until the lungs were visually free of blood. Subsequently, the lungs were removed and cut into small pieces, and the dissected tissue was then incubated in digestion solution (RPMI 1640 medium + collagenase A [5 mg/ml] and DNase I [1 mg/ml]) for 90 min at 37°C. The digested tissue was then further disrupted by gentle pipetting through a 1-ml pipette and then passed over 100-μM and 40-μM cell strainers (BD Biosciences). Finally, lung tissue digestion was stopped by adding 10 ml of RPMI 1640 medium containing 10% FCS. Macrophages contained in lung tissue homogenates were further enriched by magnetic cell separation using magnetic bead–conjugated antibodies with specificity for CD11c, known to be strongly expressed on the cell surface of lung macrophages but not PB monocytes (12). Briefly, cell suspensions were resuspended in MACS buffer (PBS/2 mM EDTA/0.5% BSA) in the presence of magnetic bead–conjugated anti-CD11c antibodies (10 μl/107 cells) for 15 min at 4°C, according to the manufacturer's instructions (Miltenyi, Bergisch-Gladbach, Germany). After incubation and washing, anti-CD11c antibody–labeled cell suspensions were passed through MS columns and CD11c-positive cells were eluted with MACS buffer, resulting in strongly enriched, highly autofluorescent CD11c-positive, CD11b-negative, and F4/80-positive lung macrophage preparations, as determined by fluorescence-activated cell sorter (FACS) analysis (data not shown).

Specificity Testing of the Employed Monoclonal Antibodies

The specificity of the employed monoclonal antibodies to selectively detect the CD45.1 or CD45.2 alloantigen on the cell surface of PB leukocytes and resident alveolar macrophages of untreated CD45.1 or CD45.2 mice was analyzed by FACS analysis. Briefly, 1 × 105 PB leukocytes or resident alveolar macrophages of either alloantigen-expressing mouse strain were placed into wells of flexible microtiter plates (BD Biosciences) and washed with PBS buffer (supplemented with 1% bovine serum albumin plus 0.2% sodium azide). Subsequently, cells were preincubated for 5 min with Fc-block (BD Biosciences) to block unspecific Fc-receptor–mediated antibody binding. Cells were then washed once in PBS buffer and incubated for 30 min with the respective PE-conjugated anti-CD45.1 or FITC-conjugated CD45.2 antibodies (1:50 dilution). Appropriately stained PB leukocyte and BAL fluid alveolar macrophages were then washed twice and FACS analysis was performed.

FACS Analysis

CD45.1 versus CD45.2 expression profiles on PB leukocytes of untreated CD45.1 or CD45.2 mice or chimeric CD45.2 mice were analyzed on a FACScan flow cytometer equipped with a 488-nm argon-ion laser and CellQuest Pro software package (BD Biosciences). For selected experiments, CD45.1 versus CD45.2 expression analysis was also done using a FACSVantage SE flow cytometer operated with a DIVA module and equipped with an argon ion laser tuned to 200 mW and a BD FACSDiva software package, as indicated. In initial experiments, PB leukocytes or resident alveolar macrophages of either mouse strain (CD45.1 versus CD45.2) were stained with either PE-conjugated CD45.1 or FITC-conjugated CD45.2 antibodies to verify the specificity of employed antibodies. In all other experiments, PB leukocytes, alveolar macrophages, or lung homogenate macrophages of the respective treatment groups were routinely stained with PE-conjugated anti-CD45.1 antibodies after appropriate gating (forward versus side scatter) of the respective cell populations. Donor cell–type specific CD45.1 PE-fluorescence emission characteristics were detected in the fluorescence two channel of the flow cytometers (F488/575), and were plotted on four (FACScan) versus five decade (FACSVantage) logarithmic histogram scales.

In selected experiments, we also analyzed induction of apoptosis in alveolar macrophages collected from LPS-treated chimeric CD45.2 mice. Cells were stained with either FITC-conjugated anti-CD45.1 antibodies for the detection of donor-type alveolar macrophages or stained with FITC-conjugated anti-CD45.2 antibodies for the detection of recipient-type alveolar macrophages for 30 min on ice followed by two washing steps, and then incubated with PE-labeled annexin V for 15 min at room temperature, according to the manufacturer's instructions (BD Biosciences). Subsequently, induction of apoptosis was analyzed after gating of the respective donor- versus recipient-type macrophages according to their forward scatter versus CD45.1 FITC or CD45.2 FITC emission characteristics, and detection of the respective PE-labeled annexin V binding was done in the fluorescence two channel of the flow cytometer.

Treatment of Mice

We established two treatment groups. In mice of treatment group one, we determined the kinetics of the constitutive alveolar macrophage turnover in unchallenged chimeric CD45.2 mice within an observation period of 1 yr (5 wk, and 2, 3, 6, and 12 mo after transplantation). In addition, chimeric CD45.2 mice of treatment group 2 were challenged with intratracheal instillations of E. coli LPS (20 μg/mouse), and the percentage of donor-type (CD45.1, recruited monocytes/macrophages) versus recipient-type (CD45.2, resident alveolar macrophages) macrophages was determined at Days 1, 3, 5, and Weeks 1, 2, 4, 8, and 12 after LPS application. Since initial experiments showed that with the chosen irradiation protocol, a maximum of > 85% CD45.1 engraftment was achieved in chimeric CD45.2 recipient mice at 5 wk after bone marrow transplant (BMT), chimeric CD45.2 recipient mice were challenged with intratracheal LPS at 5 wk after BMT. Subsequently, mice were killed with an overdose of isoflurane, and PB and BALF was collected. Determination of either donor (CD45.1) or recipient (CD45.2) phenotypes of PB leukocytes and BAL fluid macrophages of chimeric CD45.2 mice was done by FACS analysis and quantification was achieved by multiplication of the respective percentage values with absolute cell numbers.

Statistics

The data are expressed as mean ± SD. Significant differences between treatment regimen were estimated by Mann-Whitney U-test. Differences were assumed to be significant when P values were at least < 0.05.

Initial experiments were performed to verify the specificities of the chosen anti-CD45.1 versus anti-CD45.2 antibodies to detect CD45.1 versus CD45.2 alloantigen expression on PB leukocytes and alveolar macrophages. As shown in Figures 1A–1C, PB leukocytes of untreated, nontransplanted CD45.2 mice stained positive with the anti-CD45.2 antibody 104 but showed no immunoreactivity with the anti-CD45.1 antibody A20 (Figure 1C). On the other hand, PB leukocytes of untreated CD45.1 control mice did not react with the anti-CD45.2 antibody but stained strongly positive with the anti-CD45.1 antibody (Figures 1D–1F). Similarly, resident alveolar macrophages collected from untreated, nontransplanted CD45.2 mice stained strongly positive with the anti-CD45.2 antibody 104 but failed to react with the anti-CD45.1 antibody A20 (Figures 1G–1I). Conversely, resident alveolar macrophages from untreated CD45.1 mice did not stain with the anti-CD45.2 antibody 104 but showed an easily detectable, strong CD45.1 expression profile (Figures 1K–1M).

Next, we studied the kinetics of donor CD45.1 bone marrow engraftment in chimeric CD45.2 recipient mice. Figure 2 shows the analysis of donor bone marrow engraftment in recipient mice between 1 and 5 wk after BMT, and reveals that the percentage of CD45.1-positive donor-type leukocytes circulating in PB of chimeric CD45.2 mice increased in a time-dependent fashion, reaching a maximum CD45.2 to CD45.1 phenotype switching in PB leukocytes of > 85% at 5 wk after BMT (Figures 2A–2E). No further increase in engraftment efficiency was observed with the employed irradiation protocol (8+4 Gy) at time points later than 5 wk after BMT (data not shown).

Importantly, CD45.1-positive alveolar macrophages remained largely undetectable in BALF of chimeric CD45.2 mice within the first 4 wk after BMT (< 5%, Figures 3A–3C). At 5 wk after BMT, less than 10% of alveolar macrophages of untreated chimeric CD45.2 mice were of CD45.1 donor-type (Figures 3D and 4). Further analysis at 3 mo (Figures 3E–3G), 6 mo (Figures 3H–3K), and 12 mo (Figures 3L–3N) after BMT showed an alveolar macrophage turnover within the lungs of untreated chimeric CD45.2 mice of ∼ 15% (3 mo), ∼ 25% (6 mo), and ∼ 40% (12 mo) after BMT (Figures 3L–3N and 4), highlighting the slow kinetics of baseline alveolar macrophage turnover in untreated chimeric CD45.2 mice.

We next determined the alveolar macrophage turnover in chimeric CD45.2 mice receiving an intratracheal instillation of E. coli LPS at 5 wk after BMT, where the constitutive, noninflammatory alveolar macrophage turnover still ranged < 10% (Figures 3D and 4). Intratracheal application of LPS into the lungs of chimeric CD45.2 mice induced an early developing neutrophilic alveolitis peaking at Days 1–3 after LPS treatment and rapidly declining thereafter (Figure 5, upper left dot plot, and data not shown). In addition, intratracheal LPS application elicited a strong increase in total BAL fluid alveolar macrophage numbers, leading to a significant expansion of the alveolar macrophage pool at 5–7 d after LPS treatment, and declining thereafter toward baseline levels by 4 wk after treatment (Figure 6A). Importantly, FACS analysis of BAL fluid alveolar macrophages and lung homogenate macrophages of chimeric CD45.2 mice at Day 1 after LPS challenge consistently demonstrated that ∼ 15% of total macrophages were of CD45.1 donor type (Figures 6B and 6C). At 1 wk after LPS application, when the alveolar macrophage expansion reached its maximum (Figure 6A), both BAL fluid and lung homogenate macrophages were found to consist of ∼ 50% CD45.1 donor–type cells and ∼ 50% CD45.2 recipient–type cells, demonstrating that LPS-elicited alveolar macrophage expansion was based on a de novo recruitment of CD45.1 donor–type macrophages into the lungs of chimeric CD45.2 mice (Figures 6A and 6B). Importantly, FACS analysis of the alveolar macrophage and lung homogenate allotype of chimeric CD45.2 mice at later time points after LPS application showed no decrease of the CD45.1 donor–type macrophage fraction after return of total alveolar cell numbers to baseline levels, demonstrating the persistence of inflammatory elicited mononuclear phagocytes within the low-turnover resident alveolar macrophage pool. Progressive replacement of CD45.2 recipient–type by CD45.1 donor–type macrophages finally led to an exchange of the initial CD45.2-positive recipient-type alveolar macrophage pool by a > 85% CD45.1-positive donor-type alveolar macrophage pool within 12 wk after LPS application (Figures 5 and 6A–6C).

We next questioned whether both the observed turnover of CD45.2 recipient–type into CD45.1 donor–type macrophages and the subsequent decline of total BAL fluid macrophage numbers observed at later time points subsequent to LPS challenge (Figure 6A) would be mediated by an increased apoptosis in recipient and/or donor-type macrophages, ultimately contributing to alveolar macrophage homeostasis during resolving inflammation. As shown in Figure 7, under baseline conditions, alveolar macrophages from chimeric CD45.2 mice demonstrated no increased annexin V binding, thus reflecting lack of major apoptosis induction. However, starting by 1 d after endotoxin application, we found apoptosis to initially occur in CD45.2 recipient macrophages peaking at 1 wk after endotoxin challenge and declining thereafter (Figure 7A, right histograms). Importantly, we also found increased apoptosis to occur in CD45.1 donor–type alveolar macrophages, which was clearly delayed when compared with recipient-type macrophages with peak values observed at 4 until 8 wk after endotoxin application (Figures 7A and 7B). Numbers of CD45.1 versus CD45.2 macrophages exhibiting apoptosis induction rarely exceeded 10–15% of total numbers of alveolar macrophages recovered by BAL, yet these data strongly support the concept that both the observed exchange of CD45.2 into CD45.1 macrophages, and the later reduction in alveolar macrophage numbers observed during resolving inflammation, are strongly regulated by an apoptosis-dependent process.

In the present study, we employed the CD45.1 and CD45.2 alloantigen expression system in mice to generate chimeric mice expressing the CD45.1 alloantigen in PB leukocytes and the CD45.2 alloantigen in resident alveolar macrophages. In conjunction with flow cytometric analysis to differentiate donor- from recipient-type alveolar macrophages, we analyzed the extent and time course by which resident alveolar macrophages turn over under baseline compared with acute lung inflammatory conditions. We demonstrate a slow kinetic for the constitutive turnover of the resident alveolar macrophage pool in unchallenged chimeric CD45.2 mice, with an overall exchange rate of no more than ∼ 40% within an observation period of 12 mo. In contrast, intratracheal application of LPS into the lungs of chimeric CD45.2 recipient mice triggered an accelerated, increased, and apoptosis-regulated alveolar macrophage turnover, resulting in an overall > 85% turnover of recipient-type alveolar macrophages toward donor-type macrophages within 8 wk after treatment. These data are the first to demonstrate that LPS-induced acute lung inflammation triggers the robust and persistent replacement of the resident alveolar macrophage pool by newly immigrating mononuclear phagocytes.

Several radiation chimera models described in the past three decades have addressed homeostasis and pathobiology of various mononuclear phagocyte populations in mice. For example, previous experiments by Godleski and Brain using radiation chimeras bearing an antigenic specificity on the H-2 locus in conjunction with in vitro complement-mediated cytotoxicity assays showed that by Day 21 after transplantation, radiation chimeras had a > 80% replacement of recipient marrow with donor tissue (13). Another more recent study exploited ROSA 26 chimeric mice constitutively expressing β-galactosidase (β-gal) as a marker to determine the kinetics of macrophage engraftment in recipient mice. In that study, engraftment efficiencies were determined in lung sections of chimeric mice stained for both β-galactosidase activity and F4/80 monocyte/macrophage marker expression, and were reported to reach 95% in hematopoietic tissues (spleen) at 1 mo after transplantation, whereas 60% of lung macrophages were ROSA 26 positive by 12 mo after transplantation. Interestingly, at the same time, only 30% of brain microglial cells were found to be ROSA 26 positive, suggesting that the kinetics of macrophage turnover may greatly vary between different tissues (14). Finally, we recently employed green fluorescent protein (GFP) transgenic reporter mice as donors in a fetal liver transplantation (FLT) model to replace resident alveolar macrophages with a different donor genotype. We found that reconstitution of lethally irradiated recipient mice with GFP-positive donor fetal liver hematopoietic stem cells reached ∼ 85% engraftment in PB after 4 wk of FLT, whereas at the same time only a minimal alveolar macrophage turnover was observed (∼ 10%). By 10 wk after FLT, ∼ 55% of lung lavage macrophages were GFP-positive, and no significant further increase was observed at 10 mo after transplantation (15). Thus, the engraftment efficiencies of > 85% found in hematopoietic tissues of lethally irradiated CD45.2 transplant recipients in the current study are well in line with those reported in previous work, ranging between ∼ 85 and 95%.

However, greater differences among studies appear to exist for the reported baseline alveolar macrophage turnover rates, ranging between ∼ 40% (12 mo after BMT; this study), ∼ 55% (10 mo after FLT [15]), and ∼ 60% (12 mo after BMT [14]). Most likely, differences in the employed mouse systems, radiation sources, and doses applied, as well as differences between the detection systems employed to differentiate between donor and recipient cell types, may explain the reported variations in constitutive lung macrophage turnover rates. In addition, immunohistochemistry-based detection systems to enumerate ratios of recipient- to donor-type lung macrophages may possibly overestimate exchange rates within alveolar macrophages, due to the restraints in spatial resolution between vascular and interstitial versus alveolar compartments of the lung. In our hands, the currently presented radiation chimera system exploiting CD45.1 versus CD45.2 alloantigen–expressing mouse strains in conjunction with flow cytometry has the strong advantage of analyzing (1) an inflammation/activation-resistant and (2) a genetically stable and strongly expressed cell surface marker on target cells in a detection system that provides (3) a sufficiently high signal-to-noise ratio. The strong cell surface expression of CD45 on both circulating leukocytes and alveolar macrophages makes it an easily identifiable and ideal target molecule, rendering the methodology very robust and sensitive. The present system should prove useful in determining functional differences between recipient (resident)- versus donor (recruited)-type macrophages, with possible relevance for transplantation medicine, where a detailed comparative analysis of phagocytic and proinflammatory capacities between recipient-type macrophages and newly recruited donor-type macrophages residing in the same alveolar microenvironment has yet not been determined on a cellular level.

The current study presents two novel and important aspects in alveolar macrophage biology. First, it for the first time addresses the previously unappreciated alveolar macrophage turnover under acute lung inflammatory conditions: We here show that at the time of maximum alveolar macrophage expansion observed 1 wk after LPS application, the total pool of BAL fluid alveolar macrophages consisted of ∼ 50% CD45.1 donor–type and ∼ 50% CD45.2 recipient–type alveolar macrophages. At this time, the constitutive alveolar macrophage turnover in unchallenged chimeric CD45.2 mice amounted to only ∼ 10%, which indicates that alveolar macrophage expansion is virtually exclusively mediated by newly recruited donor-type macrophages. Second, the later decrease in total BAL fluid alveolar macrophage numbers observed in chimeric CD45.2 mice at > 1 wk until ∼ 8 wk during the resolving phase of inflammation was found to be due to a selective loss of CD45.2 recipient–type macrophages in the alveolar compartment, and ultimately led to a clear-cut switch from initially dominating CD45.2 recipient-type alveolar macrophages toward a > 85% CD45.1 donor–type alveolar macrophage pool after resolving inflammation. These data provide clear experimental evidence that in acutely inflamed mouse lungs, the resident alveolar macrophage pool is being replaced by newly immigrating mononuclear phagocytes. Since the chosen irradiation protocol of the current study yielded hematopoietic engraftment efficiencies of ∼ 85%, it is conceivable that the overall exchange rate of alveolar macrophages achieved under inflammatory conditions may not exceed ∼ 85%. Higher radiation doses might yield even higher hematopoietic engraftment efficiencies together with higher macrophage exchange rates within the lungs, but were not considered in the current study to avoid irradiation-induced side-effects.

Recent data in a mouse model of peritoneal inflammation suggested that in the peritoneal cavity, macrophages are cleared from the inflammatory environment by adhesion molecule–dependent mechanisms via draining lymphatics (16). In the current study, we found apoptotic events detected in both recipient and donor-type macrophages to contribute to the observed inflammation-induced macrophage turnover. Interestingly, apoptosis was initially detected in recipient-type macrophages after accelerated kinetics, whereas donor-type macrophages acquired apoptosis at much later time points. These data strongly support not only the concept that the LPS-induced exchange of recipient-type macrophages of the lung into donor-type macrophages is controlled by apoptosis, but also indicate that inflammation-induced, expanded alveolar macrophage pools of donor type are subject to apoptotic events, which may be important to regain alveolar macrophage homeostasis. Since the analysis of mediastinal lymph nodes collected from LPS-treated chimeric CD45.2 mice did not reveal elevated numbers of donor-type macrophages during resolving inflammation (data not shown), we conclude that alveolar macrophage apoptosis most probably appears to reflect the primary pathway by which alveolar macrophages are cleared from LPS-inflamed lungs.

Collectively, the present study made use of a chimeric CD45.2 alloantigen–expressing mouse model in conjunction with a FACS-based detection system to determine the alveolar macrophage turnover in mice under both baseline and acute lung inflammatory conditions. We provide evidence that under baseline conditions, alveolar macrophages are turned over with a slow kinetic not exceeding 40% after 1 yr of transplantation. In contrast, under acute lung inflammatory conditions, newly immigrated donor-type macrophages nearly completely replace the recipient resident alveolar macrophage pool within a few weeks after challenge. These data provide new insights into the pathobiology of alveolar macrophages during LPS-triggered acute lung inflammation, and may also prove useful in assigning specific functions of macrophages in acute and chronic lung infections to either the resident or recruited macrophage subset.

The authors are grateful for the excellent technical support by Regina Maus and Petra Janssen. R.M. also helped with the preparation of the FACS figures.

1. Goodman RB, Strieter RM, Martin DP, Steinberg KP, Milberg JA, Maunder RJ, Kunkel SL, Walz A, Hudson LD, Martin TR. Inflammatory cytokines in patients with persistence of the acute respiratory distress syndrome. Am J Respir Crit Care Med 1996;154:602–611.
2. Rosseau S, Hammerl P, Maus U, Walmrath HD, Schutte H, Grimminger F, Seeger W, Lohmeyer J. Phenotypic characterization of alveolar monocyte recruitment in acute respiratory distress syndrome. Am J Physiol Lung Cell Mol Physiol 2000;279:L25–L35.
3. Maus U, Huwe J, Maus R, Seeger W, Lohmeyer J. Alveolar JE/MCP-1 and endotoxin synergize to provoke lung cytokine upregulation, sequential neutrophil and monocyte influx, and vascular leakage in mice. Am J Respir Crit Care Med 2001;164:406–411.
4. Maus U, von Grote K, Kuziel WA, Mack M, Miller EJ, Cihak J, Stangassinger M, Maus R, Schlondorff D, Seeger W, et al. The role of CC chemokine receptor 2 in alveolar monocyte and neutrophil immigration in intact mice. Am J Respir Crit Care Med 2002;166:268–273.
5. Srivastava M, Jung S, Wilhelm J, Fink L, Buhling F, Welte T, Bohle RM, Seeger W, Lohmeyer J, Maus UA. The inflammatory versus constitutive trafficking of mononuclear phagocytes into the alveolar space of mice is associated with drastic changes in their gene expression profiles. J Immunol 2005;175:1884–1893.
6. Maus UA, Wellmann S, Hampl C, Kuziel WA, Srivastava M, Mack M, Everhart MB, Blackwell TS, Christman JW, Schlondorff D, et al. CCR2-positive monocytes recruited to inflamed lungs downregulate local CCL2 chemokine levels. Am J Physiol Lung Cell Mol Physiol 2005;288:L350–L358.
7. Vozzelli MA, Mason SN, Whorton MH, Auten RL Jr. Antimacrophage chemokine treatment prevents neutrophil and macrophage influx in hyperoxia-exposed newborn rat lung. Am J Physiol Lung Cell Mol Physiol 2004;286:L488–L493.
8. Maus UA, Srivastava M, Paton JC, Mack M, Everhart MB, Blackwell TS, Christman JW, Schlondorff D, Seeger W, Lohmeyer J. Pneumolysin-induced lung injury is independent of leukocyte trafficking into the alveolar space. J Immunol 2004;173:1307–1312.
9. Morse HC III. Genetic nomenclature for loci controlling surface antigens of mouse hemopoietic cells. J Immunol 1992;149:3129–3134.
10. Maus UA, Waelsch K, Kuziel WA, Delbeck T, Mack M, Blackwell TS, Christman JW, Schlondorff D, Seeger W, Lohmeyer J. Monocytes are potent facilitators of alveolar neutrophil emigration during lung inflammation: role of the CCL2–CCR2 axis. J Immunol 2003;170:3273–3278.
11. Maus U, Huwe J, Ermert L, Ermert M, Seeger W, Lohmeyer J. Molecular pathways of monocyte emigration into the alveolar air space of intact mice. Am J Respir Crit Care Med 2002;165:95–100.
12. Vermaelen K, Pauwels R. Accurate and simple discrimination of mouse pulmonary dendritic cell and macrophage populations by flow cytometry: methodology and new insights. Cytometry A. 2004;61:170–177.
13. Godleski JJ, Brain JD. The origin of alveolar macrophages in mouse radiation chimeras. J Exp Med 1972;136:630–643.
14. Kennedy DW, Abkowitz JL. Kinetics of central nervous system microglial and macrophage engraftment: analysis using a transgenic bone marrow transplantation model. Blood 1997;90:986–993.
15. Everhart MB, Han W, Parman KS, Polosukhin VV, Zeng H, Sadikot RT, Li B, Yull FE, Christman JW, Blackwell TS. Intratracheal administration of liposomal clodronate accelerates alveolar macrophage reconstitution following fetal liver transplantation. J Leukoc Biol 2005;77:173–180.
16. Bellingan GJ, Xu P, Cooksley H, Cauldwell H, Shock A, Bottoms S, Haslett C, Mutsaers SE, Laurent GJ. Adhesion molecule-dependent mechanisms regulate the rate of macrophage clearance during the resolution of peritoneal inflammation. J Exp Med 2002;196:1515–1521.
Correspondence and requests for reprints should be addressed to Ulrich A. Maus, Ph.D., Hannover School of Medicine, Laboratory for Experimental Lung Research, Feodor-Lynen-Strasse 21, Hannover 30625, Germany. E-mail:

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