In vivo and in vitro uptake of surfactant lipids by alveolar type II cells and macrophages

D. L. H. Poelma, L. J. I. Zimmermann, H. H. Scholten, B. Lachmann, J. F. van Iwaarden

Abstract

The uptake of fluorescent-labeled liposomes (with a surfactant-like composition) by alveolar macrophages and alveolar type II cells was studied using flow cytometry, in vivo by instillation of the labeled liposomes in the trachea of ventilated rats followed by isolation of the alveolar cells and determination of the cell-associated fluorescence, and in vitro by incubation of isolated alveolar cells with the fluorescent liposomes. The results show that the uptake of liposomes by the alveolar cells is time and concentration dependent. In vivo alveolar macrophages internalize more than three times as many liposomes as alveolar type II cells, whereas in vitro, the amount of internalized liposomes by these cells is approximately the same. In vitro, practically all the cells (70–75%) internalize liposomes, whereas in vivo only 30% of the alveolar type II cells ingest liposomes vs. 70% of the alveolar macrophages. These results indicate that in vivo, only a small subpopulation of alveolar type II cells is able to internalize surfactant liposomes.

  • pneumocyte
  • lung
  • liposome
  • fluorescence

pulmonary surfactant lines the alveolar capillary membrane and plays an essential role in normal lung function. It is a complex of lipids and proteins synthesized by alveolar type II cells and is stored in lamellar bodies until it is secreted into the alveolar space (4). Within the alveolus, surfactant transforms to tubular myelin, which unfolds, and the surfactant lipids are rapidly inserted in the lipid monolayer present at the air-liquid interface (4). During a breathing cycle, lipids are squeezed out of the monolayer. To maintain the primary function of the monolayer, i.e., reduction of the surface tension, the loss of lipids from the monolayer has to be compensated by renewed insertion of lipids. Because the de novo synthesis of surfactant is insufficient to correct the natural loss (i.e., inactivation of surfactant), the alveolar type II cell not only produces newly synthesized surfactant but also reutilizes inactivated surfactant, derived from the alveolar space.

The recycling of surfactant is a major pathway for surfactant in the alveolar space (14). Clearance by the mucociliary pathway (20) or degradation by alveolar type II cells and alveolar macrophages appears to be less important in the removal of surfactant lipids from the alveolar space.

One important factor in the complex system of surfactant synthesis, secretion, recycling, clearing, and degradation is the uptake of surfactant lipids by alveolar type II cells and alveolar macrophages. Numerous in vitro studies have demonstrated that both cell types can internalize surfactant lipids, although their relative contribution in the uptake of surfactant lipids in the lung remains obscure. According to Miles et al. (11), based on in vitro studies, alveolar macrophages may be responsible for all the catabolism of surfactant lipids.

In contrast, results of in vivo experiments suggest a primary role of alveolar type II cells in the uptake of surfactant lipids rather than alveolar macrophages (16, 17); however, a recent study suggests an equal contribution of both alveolar type II cells and alveolar macrophages in the uptake of surfactant lipids in vivo in the presence of surfactant protein A (SP-A) (7).

Although differences in methodology may underlie these contradictory findings, another explanation may be the difference between in vivo and in vitro experiments. Study of the uptake of lipids by alveolar cells in vivo and in vitro, using a similar technique, has the advantage that the contribution of the alveolar type II cells and alveolar macrophages in the uptake of lipids in the lung can be assessed, as can the influence of cell isolation and environmental factors. In the present study, we have studied the uptake of fluorescent-labeled liposomes by alveolar type II cells and alveolar macrophages in vivo as well as in vitro using flow cytometry.

In contrast to most studies on the uptake of surfactant lipids by type II cells or alveolar macrophages, which focus on its main component, saturated phosphatidylcholine (PC; i.e., dipalmitoyl phosphatidylcholine; DPPC) (16-18), in the present study, we used fluorescent liposomes with a composition similar to natural surfactant: DPPC, PC, phosphatidylglycerol (PG), phosphatidylinositol (PI), phosphatidylethanolamine (PE), and cholesterol (weight ratio 55:21:8:2:6:8) (20).

An advantage of using flow cytometry to study the uptake of lipids by alveolar type II cells and alveolar macrophages is that it allows us to determine whether all the cells are able to internalize lipids or only subpopulations. To date, it is not known whether all alveolar macrophages and/or alveolar type II cells are involved in the uptake of lipids. We have developed a new method to study the uptake of surfactant lipids by alveolar type II cells and alveolar macrophages in vivo as well as in vitro using fluorescent-labeled surfactant-like liposomes.

MATERIALS AND METHODS

Ethical guidelines.

This study was approved by the Institutional Animal Committee at the Erasmus University Rotterdam.

Liposome preparation.

To prepare surfactant-like liposomes, the following lipids were mixed: DPPC, PC, PG, PI, rhodamine-labeled PE [rhodamine 1,2 dihexadecanoyl-sn-glycero-3-phosphoethanolamine (DHPE); Molecular Probes, Leiden, The Netherlands], and cholesterol in a weight ratio of 55:21:8:2:6:8 and dried under a stream of nitrogen gas. The lipids were purchased from Sigma (Zwijndrecht, The Netherlands), unless stated otherwise. The liposomes were suspended in PBS at a concentration of 1 mg of lipids/ml using glass pearls and vortexing. Immediately before use, the liposome suspension was sonicated for 2 min on ice using an ultrasonic disintegrator (Branson Sonifier 250) to prepare small unilamellar liposomes. These liposomes were used unless stated otherwise. Large multilamellar liposomes were prepared by sonicating the liposome suspension in a water bath sonicator (Branson 5210) at 37°C for 3 min. The size of the liposomes was determined by dynamic light scattering at 25°C with a Malvern 4700 system using a 25-mW He-Ne laser (NEC, Tokyo, Japan) and the automeasure version 3.2 software (Malvern). As a measure of particle size distribution of the dispersion, the system reports a polydispersity index (pd). This index ranges from 0.0 for a monodisperse and up to 1.0 for an entirely polydisperse dispersion. The “small” liposomes had a size of 161 nm pd 0.35, and the “large” liposomes had a size of 1,455 nm pd 0.65, significantly larger than the small liposomes.

Intratracheal instillation of fluorescent liposomes.

The studies were performed with male Sprague-Dawley rats (IFFA Credo) with a body wt of 314 ± 18 g. After induction of anesthesia with a mixture of nitrous oxide (66%), oxygen (33%), and isoflurane (1–2%), a sterile polyethylene catheter (0.8-mm outer diameter) was inserted into one of the carotid arteries. The animals were then tracheotomized, and a sterile metal cannula was inserted into the trachea.

After these surgical procedures, gaseous anesthesia was ended and replaced with an intraperitoneal injection of pentobarbital sodium (60 mg/ml of Nembutal; Algin BV, Maassluis, The Netherlands) at a dose of 30 mg/kg body wt every hour.

Muscle relaxation was induced and maintained by an hourly intramuscular injection of pancuronium bromide (2 mg/kg of Pavulon; Organon Technika, Boxtel, The Netherlands). The animals were then mechanically ventilated with a Servo ventilator 300 (Siemens-Elema, Solna, Sweden) set to pressure control mode using a frequency of 30/min, an inspiratory-expiratory ratio of 1:2, a positive end-expiratory pressure (PEEP) of 2 cmH2O, a peak inspiratory pressure (PIP) of 12 cmH2O, and Fi O2 was set to 1.

Before instillation of the labeled liposomes, PEEP was increased to 6 cmH2O, and PIP was increased to 26 cmH2O. After disconnection from the ventilator, the liposomes were administered intratracheally at the indicated doses. The suspension of liposomes (1 mg of lipids/ml, unless stated otherwise) was administered as a bolus of 3 ml/kg, followed by a bolus of air (12 ml/kg) directly into the endotracheal tube via a syringe, and the animals were immediately reconnected to the ventilator. Thirty minutes after instillation of the liposomes, PEEP was reduced to 2 cmH2O and PIP to 12 cmH2O.

Arterial blood gases were measured with conventional methods (ABL 505; Radiometer, Copenhagen, Denmark) at the start of ventilation, immediately after instillation of the liposomes, and every 30 min thereafter. One hour after ventilation (unless stated otherwise), the animals were killed by exsanguination via the abdominal aorta, and the alveolar cells were isolated to determine the cell-associated fluorescence. Control animals were killed immediately after anesthesia, and their isolated alveolar type II cells and alveolar macrophages were used to correct for autofluorescence.

Instillation of a large volume of liposomes.

After the surgical procedures, one group of animals was instilled with a large volume of fluorescent-labeled liposomes (30 ml/kg; 1 mg of lipids/ml). The liposomes were retrieved from the lungs immediately after instillation (recovery 90%). One hour after instillation, the animals were killed, alveolar type II cells and alveolar macrophages were isolated, and cell-associated fluorescence was determined.

Isolation of alveolar type II cells and alveolar macrophages.

Before isolation of the cells, the thorax was opened, and the blood cells were removed from the lungs by perfusing the pulmonary artery with saline (37°C) supplemented with 20 IU of heparin (Leo Pharma, Weesp, The Netherlands). The lungs were removed from the thoracic cavity en bloc and lavaged with 10 ml of solution 1(140 mM NaCl, 5 mM KCl, 2.5 mM Na2HPO4 · 2H2O, 10 mM HEPES, 6 mM glucose, and 0.2 mM EGTA, pH 7.40) at 22°C. This procedure was repeated four times. The lung lavages were pooled per animal and centrifuged (100 g, 10 min, 4°C). The cellular pellet, i.e., alveolar macrophages, was suspended in solution 2 (140 mM NaCl, 5 mM KCl, 2.5 mM phosphate buffer, 10 mM HEPES, 6 mM glucose, 2.0 mM CaCl2, and 1.3 mM MgSO4) to a concentration of 2 × 106 cells/ml and stored on ice until further use. The alveolar type II cells were isolated according to Dobbs et al. (2). Alveolar type II cells were suspended in solution 2 at a concentration of 2 × 106 cells/ml and stored on ice until further use. Alveolar macrophages were identified using monoclonal antibodies specific for rat macrophages (ED9), and alveolar type II cells were identified using an alkaline phosphatase assay as described by Edelson et al. (3). The average yield of alveolar type II cells was 16 × 106 and 5 × 106 alveolar macrophages/rat.

In vitro assay to determine uptake of fluorescent-labeled liposomes.

Alveolar type II cells and alveolar macrophages were isolated from control animals as described above and were suspended in solution 2 to a concentration of 2 × 106 cells/ml. A total of 3 × 105 cells was incubated with various concentrations of liposomes at 37°C (final vol 500 μl) in a shaking water bath. After 1 h, the incubation was terminated by addition of 2 ml of ice-cold PBS. The cell suspension was centrifuged at 100g for 10 min at 4°C. The supernatant was removed, and the cells were suspended in 2 ml of ice-cold PBS and centrifuged again. This wash procedure was repeated twice. Finally, the pellet was resuspended in 200 μl of cold PBS, and cell-associated fluorescence was determined as described below.

Flow cytometry.

Cell-associated fluorescence of the alveolar type II cells and alveolar macrophages as a measure for internalized liposomes was determined using flow cytometry (FACSCalibur; Becton Dickinson). The cell-associated fluorescence of 15,000 cells was determined. Alveolar macrophages and alveolar type II cells derived from control animals were used in each experiment to determine the autofluorescence of the cells. Subsequently, the mean cell-associated fluorescence was determined only for those cells that had a higher fluorescence than that caused by autofluorescence (gated cells). For the cell-sorting experiment, the FACSCalibur was set to the sorting mode, and only those cells with a mean fluorescence higher than the autofluorescence were collected and analyzed.

Localization of cell-associated fluorescence.

To inspect the localization of the cell-associated fluorescence, confocal micrographs of alveolar cells were obtained using a confocal microscope (Zeiss LSM 410). Images were created with standard objectives and photomultiplier tubes dedicated to the appropriate excitation and emission spectra of rhodamine (excitation 541 nm and emission 572 nm). Images of alveolar cells were serial sectioned at a depth of 0.5 μm to distinguish cell membrane-associated fluorescence from true intracellular fluorescence.

Statistical analysis.

Differences in blood gas values over time were analyzed using a repeated-measurement ANOVA followed by an independentt-test. Differences between animals that received fluorescent-labeled liposomes and the control group were determined with an independent sample t-test. Differences between the animals receiving a normal volume of liposomes and the animals receiving a high volume of liposomes were analyzed using an independent t-test. Differences were considered statistically significant at P < 0.05. Values are expressed as means ± SE.

RESULTS

Isolated alveolar type II cells and alveolar macrophages.

The forward and sideward scatter plots in Fig.1 demonstrate that alveolar type II cells and alveolar macrophages isolated from controls consist of heterogeneous cell populations. The cell-associated fluorescence of alveolar cells isolated from rats 1 h after intratracheal instillation of fluorescent-labeled liposomes was clearly higher than the control cells. The cell-associated fluorescence of the alveolar macrophages is higher than the fluorescence of the alveolar type II cells, suggesting more uptake of fluorescent liposomes by alveolar macrophages. Instillation of the fluorescent-labeled liposomes had no significant effect on oxygenation levels of the animals during ventilation (results not shown).

Fig. 1.

Flow cytometry of alveolar cells. Alveolar type II cells (A, C, E) and alveolar macrophages (B, D, F) were isolated from control animals (A–D) and from animals 1 h after intratracheal instillation of fluorescent liposomes (E andF). The forward and sideward scatter (SSC) plots demonstrate the presence of heterogeneous populations of alveolar type II cells (A) and alveolar macrophages (B). Autofluorescence of control alveolar type II cells (C) and control alveolar macrophages (D) was determined. Total cell-associated fluorescence (FL2-PI) was determined of alveolar type II cells (E) and alveolar macrophages (F) isolated from animals that were intratracheally instilled with fluorescent liposomes. Marker M1 (C–F) indicates the mean cell-associated fluorescence of the cells that have internalized the fluorescent liposomes (gated cells).

In vivo uptake of fluorescent liposomes by alveolar cells.

To characterize the in vivo uptake of fluorescent-labeled liposomes by alveolar cells, the cells were isolated at different time points (0.5–3 h) after instillation of the fluorescent-labeled liposomes. The cell-associated fluorescence of alveolar type II cells and of alveolar macrophages demonstrated a time-dependent increase reaching a plateau in 1 h (Fig. 2). In addition, concentration dependency was determined by instilling fluorescent-labeled liposomes of different concentrations (0.5, 1, 2, 3, and 5 mg/ml) and isolating the alveolar cells 1 h after instillation (Fig. 3). At a concentration of 1 mg/ml, the cell-associated fluorescence of the alveolar type II cells was half-maximal, whereas for alveolar macrophages, the half-maximal uptake was observed at an intratracheal dose of 2 mg/ml of fluorescent liposomes. The percentage of alveolar cells involved in the uptake of fluorescent liposomes (gated cells) reached a maximum at a concentration of 1 mg/ml (Fig. 3). Approximately 30% of the alveolar type II cells participated in the uptake of fluorescent-labeled liposomes. In contrast, ∼70% of the alveolar macrophages internalized the fluorescent-labeled liposomes (Fig. 3). At an intratracheal dose of 3 mg of fluorescent liposomes, the uptake by the alveolar cells was maximal. The mean cell-associated fluorescence of the alveolar macrophages involved in the uptake was 3.26 ± 1.24 times higher than that of the alveolar type II cells; this indicates 3.26 times more uptake of liposomes by alveolar macrophages than alveolar type II cells.

Fig. 2.

Time-dependent uptake of fluorescent liposomes. At every time point (30 min, 1, 2, and 3 h after instillation of the liposomes), 5 animals were killed, alveolar type II cells (A) and macrophages (B) were isolated, and mean cell-associated fluorescence was determined. At all time points, fluorescence differed significantly from the control group (values are means ± SE).

Fig. 3.

Concentration-dependent uptake of liposomes in vivo. One hour after instillation of the indicated concentrations of fluorescent liposomes, the alveolar cells were isolated. The mean cell-associated fluorescence of the gated alveolar type II cells (A) and alveolar macrophages (C) was determined. In addition, the percentage of gated alveolar type II cells (B) and alveolar macrophages (D) was determined. At all concentrations, fluorescence and percentage of gated cells differed significantly from the control groups (n = 4 rats at every concentration; values are means ± SE).

Uptake of unilamellar vs. multilamellar liposomes.

As demonstrated by Griese and Reinhardt (6), smaller-sized particles are preferentially taken up by the alveolar type II cells. We studied the effect of the size of the labeled liposomes on the uptake in vivo by instilling either small unilamellar liposomes, created by ultrasonification, or large multilamellar liposomes, sonicated in a warm water bath sonicator (19). No differences were observed in the uptake of both types of liposomes, unilamellar or multilamellar (Table 1).

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Table 1.

Size-dependent uptake of fluorescent liposomes

Localization of the cell-associated fluorescence.

Confocal laser microscopy was used to ascertain that the measured cell-associated fluorescence was caused by intracellular presence of fluorescent-labeled liposomes. Confocal scans through the middle of the cell (Fig. 4) show that the fluorescence is not limited to the circumference of the cell and show a punctuate fluorescence throughout the cell except for the nucleus.

Fig. 4.

Confocal microscopic images of alveolar type II cells and alveolar macrophages. Alveolar type II cells (A) and alveolar macrophages (B) isolated 1 h after instillation of fluorescent-labeled liposomes were mounted on glass coverslips. Confocal laser microscopy demonstrated a punctuate distribution of fluorescent label not limited to the cell surface, which did not localize to the nuclei, indicating internalization of the fluorescent liposomes by both cell types. Because untreated control cells did not show this fluorescence at the same microscope settings, we conclude that the observed internal fluorescence of treated cells was due to the uptake of fluorescent-labeled liposomes. Bar, 5 μm.

Identification of cells that internalize the fluorescent liposomes.

We found that in vivo only 30% of the type II cells isolated from rats intratracheally instilled with fluorescent liposomes were involved in the uptake of fluorescent liposomes. The average percentage of cells identified as alveolar type II cells in the alveolar type II cell isolate was ∼80%. Theoretically, it is possible that a substantial number of the cells that internalize fluorescent liposomes in alveolar type II cell isolate are not type II cells but contaminating cells such as macrophages, etc. To exclude this possibility, rats were intratracheally instilled with fluorescent liposomes. The alveolar type II cells and alveolar macrophages were isolated. The alveolar cells with a mean fluorescence higher than the autofluorescence of the control cells were sorted and collected. As shown in Table2, 80 ± 5% (n = 3) of these fluorescent cells present in the alveolar type II cell isolate were identified as alveolar type II cells using alkaline phosphatase assay, whereas 92 ± 5% (n = 3) of the alveolar macrophages could be identified as macrophages, as was demonstrated using a monoclonal antiserum directed against macrophages (ED9).

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Table 2.

Identification of fluorescent cells

Distribution of the liposomes.

To determine whether the low number of alveolar type II cells internalizing the liposomes was due to an inhomogeneous distribution of liposomes in the lungs, one group of animals was instilled with a high volume of fluorescent-labeled liposomes.

Instillation of a large volume of liposomes, which was immediately redrawn from the lungs, did not result in a significant difference in the percentage of cells responsible for the fluorescence, nor was there any significant difference in the level of fluorescence between the animals receiving the liposomes as a bolus and the animals receiving a high volume of liposomes (Table 3). Interestingly, the number of alveolar macrophages involved in the uptake of liposomes was significantly lower in the high volume group than in the low volume, standardly used group.

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Table 3.

Distribution of fluorescent liposomes

In vitro measurement of uptake.

In vitro experiments with different concentrations of fluorescent-labeled liposomes demonstrated an apparent maximal uptake at 20 μg/ml for both cell types (Fig.5). The percentage alveolar type II cells responsible for the uptake was maximal at 100 μg/ml at 67.6 ± 11.1%. The percentage of alveolar macrophages reaches its maximum of 73 ± 34.7% at 20 μg/ml (Fig. 5). The mean cell-associated fluorescence did not significantly differ between the alveolar type II cells and the alveolar macrophages.

Fig. 5.

Concentration-dependent uptake of liposomes in vitro. Alveolar cells were isolated from control animals and incubated for 1 h at 37°C with the indicated concentrations of fluorescent liposomes. The mean cell-associated fluorescence of gated alveolar type II cells (A) and alveolar macrophages (C) was determined. In addition, the percentage of gated alveolar type II cells (B) and alveolar macrophages (D) was determined. At all concentrations, fluorescence and percentage gated cells differed significantly from the control group (n = 4 incubations at every concentration; values are means ± SE).

The mean fluorescence of the alveolar macrophages in vitro is 5.12 ± 1.01 times lower compared with the fluorescence of the alveolar macrophages in vivo.

To get an impression of the amount of lipids that are only bound to the cellular membranes of the cells, we incubated the cells on ice with the fluorescent liposomes for 1 h. For alveolar type II cells, 12 ± 4% (n = 3) of the total cell population had a mean fluorescence [28 ± 7 (n = 3)] higher than the autofluorescence. For alveolar macrophages, 18 ± 3% (n = 3) of the cells had a mean fluorescence [36 ± 5 (n = 3)] higher than the autofluorescence.

DISCUSSION

In the present study, we used fluorescence-labeled liposomes to study the uptake of lipids by alveolar cells. For the in vivo experiments, these liposomes were intratracheally instilled in mechanically ventilated rats. The choice to mechanically ventilate the animals was made on the basis of ensuring that the liposomes are instilled in a completely opened lung rather than in a lung in which atelectasis is present. The ventilator settings used, peak of 26 cmH2O and a PEEP of 6 cmH2O, resulted in a mean airway pressure of 13 cmH2O. It has been demonstrated that ventilating with mean airway pressure up to 15 cmH2O does not affect the surfactant metabolism and does not induce lung injury (8).

By using different concentrations, we have demonstrated a concentration-dependent uptake, reaching a maximum at 3 mg/ml. A time-dependent uptake was demonstrated by isolating the alveolar cells at different time points after instillation of the liposomes, similar to previous studies using radioactive-labeled liposomes (5, 13,17). The use of a composition similar to natural surfactant diminishes the effects of single components and allows the study of uptake of surfactant as a whole complex of different lipids.

The use of confocal laser microscopy allowed us to demonstrate the intracellular presence of the fluorescent-labeled liposomes. The cell-associated fluorescence was, therefore, caused by internalized liposomes rather than by liposomes associated with the cell membrane of the alveolar cells.

The use of fluorescence enabled us to quantitate the fluorescence per individual cell, which led to intriguing results. Interestingly, only 30% of the alveolar type II cells is involved in the uptake of liposomes. After instillation of liposomes with different concentrations, the number of cells taking part in the uptake of the liposomes does not increase above ± 30%. Identification of the isolated cells demonstrated a similar amount of type II cells in the type II cell isolate, as reported in other studies (12, 16,17). The cells with a fluorescence higher than the autofluorescence were identified as type II cells, demonstrating that the measured cell-associated fluorescence was indeed caused by type II cells. The low number of alveolar type II cells involved in the uptake of lipids in vivo is intriguing because the in vitro experiments demonstrated a much larger number of type II cells (70%) participating in the uptake. Instillation of a high volume of labeled liposomes, distributing the liposomes throughout the lung, did not result in an increase in type II cells taking up the liposomes. Hence, the low number of participating type II cells in vivo is not caused by an insufficient distribution or by cells other than alveolar type II cells in the cell isolate. In other words, the difference is either caused by the isolation procedure or is due to the absence of environmental influences in vitro. It seems unlikely that the isolation procedure stimulates the alveolar type II cells to take up lipids. It has been observed that a stressful isolation procedure can cause secretion of lamellar bodies (21). In addition, the turnover, degradation, and resecretion continues during the 1-h incubation. Therefore, the measured fluorescence in vivo is an underestimation of the actual uptake of the fluorescence-labeled liposomes. On the other hand, the fluorescence per cell is only slightly increased in vitro, whereas the number of cells participating in the uptake is significantly increased. Therefore, these results suggest a possible regulating effect of the alveolar environment on the uptake of lipids by alveolar type II cells as well as the presence of a subpopulation of alveolar type II cells. This subpopulation of alveolar type II cells is activated either during the isolation procedure or by the absence of the environmental factors, or degradates/resecretes the fluorescent-labeled liposomes more rapidly. Alveolar type II cells have been shown to interact with alveolar macrophages (9,10, 15), although it remains unclear if these cells regulate each other's uptake of lipids.

In addition, an important role of the environment is also seen in the uptake of lipids by alveolar macrophages. Both in vivo and in vitro, the same number of alveolar macrophages takes part in the uptake (70%), although most interestingly, the alveolar macrophages take up more lipids in vivo than in vitro. The instillation of liposomes with different concentrations demonstrated that the uptake of liposomes by alveolar macrophages reaches a plateau phase, both in vivo and in vitro. When comparing the mean fluorescence per cell at this plateau, it is obvious that in vivo, the alveolar macrophages take up approximately five times as many liposomes than in vitro. In other words, both the alveolar type II cells and alveolar macrophages interact with the environment.

Additionally, the results of the instillation of a high volume of liposomes suggest the presence of subpopulations within the alveolar macrophages. When a high volume of liposomes (30 ml/kg) is instilled and removed from the lung, the same number of alveolar macrophages is removed as in a lung lavage. One hour after instillation and withdrawal of the high volume, the alveolar cells were isolated, and cell-associated fluorescence was determined. The alveolar type II cells showed no significant difference compared with the cells derived from animals instilled with the normal volume (1 ml). In contrast, the alveolar macrophages showed a significant decrease in the number of cells taking up the liposomes. The mean fluorescence per cell did not differ significantly, suggesting that a part of the cells that is involved in the uptake of fluorescent liposomes was removed by the instillation and withdrawal of a high volume of liposomes. In this primary “lavage,” the most mobile cells are removed, whereas the more resident cells are not likely to be removed. Although there might be influx of new alveolar macrophages by chemotaxis, their contribution in the clearance of the liposomes from the alveolar space is time dependent, as demonstrated in the present study. We therefore conclude that the relatively short time of exposure to the liposomes of recently influxed cells can explain the minor contribution of these newly influxed cells. The mean fluorescence per cell did not differ, indicating that the cells were not negatively influenced by this lavage, although the low number of cells involved suggest that the liposomes are preferentially taken up by the mobile macrophages rather than by the resident, adherent cells.

To date, their relative contribution is under discussion. Some studies indicate a minor role of the alveolar macrophages in the uptake of lipids (17). Others indicate a larger contribution of these cells to the clearance of surfactant (7, 11). We have found, when comparing the alveolar type II cells with the alveolar macrophages, that the alveolar macrophages take up approximately two to three times more liposomes. When correcting for the number of cells within the lung, it is clear that the alveolar type II cells contribute ± 60–70% of the clearance of surfactant from the alveolar space. There are about 10–15 times as many type II cells than macrophages (calculations using Ref. 1), and although only 30% of the type II cells are involved in the clearance of the liposomes compared with 70% of the alveolar macrophages, the alveolar type II cells still outnumber macrophages five to seven times with regard to the number of cells involved in the clearance of liposomes. The macrophages take up two to three times more liposomes than the alveolar type II cells. Therefore, the alveolar type II cells contribute about two to three times more than the alveolar macrophages.

In the present study, we used fluorescence-labeled liposomes with a composition similar to that of natural surfactant without surfactant proteins. Although studies in the past have demonstrated a possible role of the apoproteins, especially SP-A, on the uptake of surfactant by both alveolar type II cells and alveolar macrophages (7), the present study focuses on the method of measuring the uptake rather than studying the role of the apoproteins. The absence of the surfactant proteins might account for the different findings in the contribution of alveolar macrophages in the present study, although the interspecies variation is another option.

In summary, using fluorescence-labeled liposomes, we have demonstrated an important role of the environment on the uptake of surfactant liposomes by alveolar cells as well as the presence of possible subpopulations of alveolar type II cells and macrophages involved in the uptake.

Acknowledgments

The authors thank Dr. G. Kraal, Dept. of Cell Biology and Immunology, Free University, Amsterdam, The Netherlands, for supplying ED9; Dr. A. B. Houtsmuller, Dept. of Pathology and Josephine Nefkens Institute, Erasmus University Rotterdam, The Netherlands, for confocal microscopy imaging; L. van Bloois, Department of Pharmaceutics, Faculty of Pharmacy Utrecht, The Netherlands; S. C. Estevão and G. Schoonhoven for technical assistance; and Laraine Visser-Isles for English language editing.

Footnotes

  • This study was supported by LEO Pharmaceutical Products, Ballerup, Denmark.

  • Address for reprint requests and other correspondence: J. F. van Iwaarden, Laboratory of Pediatrics, Erasmus Univ. Rotterdam, PO Box 1738, 3000 DR Rotterdam, The Netherlands (E-mail:vaniwaarden{at}kgk.fgg.eur.nl).

  • The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

  • May 3, 2002;10.1152/ajplung.00478.2001

REFERENCES

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