ARTICLE |
Correspondence to: James M. Anderson, Inst. of Pathology, Case Western Reserve University, Cleveland, OH 44106..
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Summary |
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During the inflammatory response to an implanted biomaterial, monocytes undergo a striking phenotypic progression of differentiation into macrophages, which may subsequently fuse to form foreign body giant cells (FBGCs). Taking advantage of an in vitro system of cytokine-induced FBGC formation together with the optical slicing capabilities of a confocal microscope, we investigated the cytoskeletal reorganization and adhesive structure development during this dramatic morphological progression. Human monocytes demonstrated diffuse cytoplasmic staining of adhesive structural proteins. Punctate filamentous (F)-actin structures appeared along the ventral cell membrane of macrophages and were identified as the core of podosome adhesive structures by the distinctive ring staining of vinculin, talin, and paxillin around the F-actin. Cytokine-induced FBGCs were characterized by a restriction of podosomes to the extreme periphery of the ventral cell surface. Although macrophages and FBGC contained equivalent amounts of F-actin, significantly more F-actin was located within 1 µm of the ventral plasma membrane in FBGCs compared to macrophages. Taken together, these results provide new information on the dynamic cytoskeletal reorganization and adhesive structure development that occur during phenotypic progression from human monocytes to macrophages to FBGC. Furthermore, they suggest the acquisition of functional specializations on FBGC formation, which may enhance our understanding of chronic inflammatory processes. (J Histochem Cytochem 47:6574, 1999)
Key Words: macrophage, foreign body giant cell, F-actin, podosomes, chronic inflammation, confocal microscopy
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Introduction |
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Monocyte-derived macrophages are extremely versatile cells which are believed to be critical mediators of the host reaction to biomedical material implants (
Cytoskeletal participation has been demonstrated for many monocyte/macrophage functions that are critical during the inflammatory response, including migration (
Interestingly, several different types of adhesive structures (
Recent advances in confocal fluorescence microscopy, principally its capability for qualitative and quantitative evaluation of three-dimensional organization, now facilitate examination of cytoskeletal reorganization and adhesive structure development during the remarkable phenotypic progression from monocytes to macrophages and particularly during macrophagemacrophage fusion to form FBGCs. Using an in vitro system of cytokine-induced human macrophage fusion and FBGC formation (
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Materials and Methods |
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Monocyte Isolation and Culture
Human blood monocytes were isolated from the venous blood of unmedicated donors by a nonadherent density centrifugation method as described (
On Days 3 and 7 of incubation, the medium was replaced with 25% heat-treated (56C water bath for 1 hr) autologous serum in RPMI and 10 ng/ml interleukin-13 (IL-13; R & D Systems, Minneapolis, MN) was added to induce macrophage fusion and FBGC formation as indicated. For microfilament disruption experiments, 5 µM cytochalasin D (Sigma Chemical; St Louis, MO) was added to Day 10 cultures for 2 hr before fixation. Samples were fixed by rinsing twice for 5 min each with warmed PBS and covering with 3.7% formaldehyde or 3.7% formaldehyde containing 0.5 µM taxol (Sigma) in PBS for 20 min. After three additional rinses with PBS, cells were permeabilized with 0.2% Triton X-100 (Sigma) in PBS for 5 min.
Cytoskeletal and Cytoplasmic Protein Fluorescence Staining
Confocal scanning laser microscopy (MRC-600; Bio-Rad, Hercules, CA) was used to image cells with double labels of rhodaminephalloidin for F-actin and an indirect immunofluorescence tag for another cytoskeletal or cytoplasmic protein. Nonspecific sites were blocked with a 1:100 dilution of goat serum (DAKO; Carpinteria, CA) in PBS for 30 min at 37C. Blocking serum was removed and the primary antibody solutions were incubated with cells for 1 hr at 37C. Primary antibodies were diluted in PBS containing 3% BSA (Sigma) as follows: anti-tubulin 1:50 (Boehringer Mannheim; Indianapolis, IN); anti-vimentin 1:10 (Boehringer Mannheim); anti-talin 1:100 (Serotec; Oxford, UK); anti-vinculin 1:15 (Serotec); anti-paxillin 1:200 (Transduction Laboratories; Lexington, KY); anti-gelsolin, 1:500 (Sigma); and anti-focal adhesion kinase (FAK) 1:50 (Santa Cruz Biotechnology; Santa Cruz, CA). Anti-L-plastin antibodies were the generous gift of Dr. Samuel L. Jones (Washington University; St Louis, MO). Each monoclonal primary antibody had an isotype-matched control IgG (DAKO) and each polyclonal antibody had a species-matched nonspecific control IgG (DAKO) that was run in parallel. After the primary antibody incubations, the cultures were rinsed four times for 5 min each with PBS. Rhodaminephalloidin (Molecular Probes; Eugene, OR) at a final dilution of 1:150 was added to BODIPY FL-conjugated goat anti-mouse IgG (Molecular Probes) that was diluted 1:100 in PBS. The secondary antibody solution was incubated with cells for 30 min at room temperature. After three additional 10-min washes in PBS, samples were removed from the culture plates and mounted on glass slides with Gel/Mount (Biomeda; Foster City, CA).
For confocal imaging, control stains were set to a black background. Positive samples were then viewed at the same laser intensity as well as aperture, gain, and blacklevel settings. Optical slices approximately 1 µm thick were taken for each sample, beginning at the coverslip surface and ending at the apical surface of the adherent cell. Presented images of cellular cytoskeletal organization are projections of all of the slices from a sample. Images depicting adhesive structure organization are single optical slices taken from the basal cell surface.
Quantitation of Cellular Fluorescence
Samples that were double labeled for F-actin and gelsolin were used for quantitative measurements. Confocal optical slices encompassing the entire volume of individual cells were isolated using three-dimensional reconstruction software (microVoxel; Indec Systems, Capitola, CA). The fluorescence intensity of F-actin from the entire cell was measured to compare the total F-actin content in macrophages and FBGCs. F-actin participation in adhesive structures was quantified by measuring the fluorescence intensity of F-actin that co-localized with gelsolin at the ventral cell surface of macrophages and FBGCs and expressing the results as a percentage of the total cellular F-actin. Sixty macrophages and 60 FBGCs from four different donors were measured, and the unpaired Student's t-test was used for statistical analysis (StatView; Abacus Concepts, Berkeley, CA).
Measurement of Cell Diameter
Cells were stained sequentially with MayGrünwald stain (Sigma) for 1 min, phosphate buffer (pH 7.4; Sigma) for 1 min, Giemsa stain (
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Results |
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Dynamic F-Actin Reorganization Accompanies Monocyte-to-Macrophage Morphological Progression
To provide a context in which to understand macrophage to FBGC cytoskeletal reorganization, we first sought to describe the monocyte-to-macrophage cytoskeletal changes occurring during a 10-day culture period.
Two hours after freshly isolated monocytes were plated, they appeared fairly homogeneous as judged by phase-contrast microscopy. Most monocytes displayed a rounded morphology (mean diameter of 12.4 ± 0.1 µm) with characteristic kidney-shaped nuclei that occupied the majority of the cell volume (not shown). By confocal imaging, F-actin was observed to surround the nucleus in a diffuse pattern with concentrations in membrane ruffles, delineating the cell boundaries (Figure 1A). Some monocytes exhibited an elongated morphology suggestive of motility.
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By 3 days of culture, monocyte/macrophage F-actin redistributed into intensely staining punctate foci that were restricted to the substrate-attached side of adherent cells (Figure 1B). Punctate F-actin was visible across the entire ventral cell surface in spread cells, whereas it was restricted to the lamellipodia and uropods of elongated cells. In monocytes/macrophages with either morphology, microfilaments were visible along the plasma membrane throughout the volume of the cell.
In the absence of added cytokine, most cells acquired a spread morphology by Day 10 of culture (Figure 1C). These macrophages resembled those at Day 3 but were larger in diameter (25.6 ± 0.57 µm).
Cytoplasmic Proteins Associate with Punctate Filamentous Actin to Form Podosomes
The punctate F-actin fluorescence along the ventral cell membrane is indicative of adhesive structure formation at these sites (
Freshly isolated and cultured monocytes did not form prominent ventrally located punctate actin structures after 2 hr of culture. Likewise, cytoplasmic proteins known to participate in the formation of adhesive structures, i.e., vinculin, talin, and paxillin (
On development of punctate F-actin structures, vinculin, talin, and paxillin formed ring-like structures around the F-actin but did not co-localize with it, as demonstrated by paxillin in Figure 2A. Although vinculin and F-actin are considered hallmarks of focal contacts (
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The lack of co-localization between F-actin and these structural proteins precluded their identification as focal contacts or close contacts. However, the arrangement of vinculin, talin, and paxillin in ring structures surrounding F-actin cores is consistent with podosome adhesive structures (
The presence of paxillin prompted investigation for the presence of FAK, the activation of which has been described as an important step in the recruitment of paxillin to developing adhesive structures in rat embryo fibroblasts and mouse 3T3 cells (
Induction of Macrophage Fusion to Form FBGCs Results in Further Cytoskeletal and Adhesive Structure Polarizations
Because IL-13 has been demonstrated to induce macrophage fusion and FBGC formation (
In contrast to macrophages, punctate F-actin at the ventral cell surface was restricted to the extreme periphery of FBGCs and was very dense (Figure 3A). FBGCs contained dense meshworks of microtubules that usually formed circles concentric to the plasma membrane (Figure 3B). Microtubules were present in the FBGC periphery but did not terminate in the podosomes where punctate F-actin structures were concentrated. Furthermore, microtubules and intermediate filaments appeared to encase individual nuclei in FBGCs. Very dense meshworks of intermediate filaments radiated throughout the cytoplasm and terminated in fibers along the plasma membrane (Figure 3C).
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Vinculin, talin, and paxillin localized around the punctate F-actin, forming podosomes, although distinct podosomes were not always visible because of the dense and intense F-actin staining (not shown; and Figure 2C). Gelsolin remained co-localized with punctate F-actin in FBGC (Figure 2D).
To determine whether F-actin content and distribution were quantitatively different between macrophages and the morphologically distinct FBGCs, rhodaminephalloidin-labeled F-actin fluorescence intensity was measured (Table 1). Although total F-actin, normalized for volume, was equivalent between macrophages and FBGCs, significantly more F-actin was located within 1 µm of the culture surface in FBGCs.
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F-actin participation in podosome structures was quantified by measuring the fluorescence intensity of F-actin that co-localized with gelsolin at the ventral cell surface of macrophages and FBGCs (Table 1). Although the majority of the total F-actin in macrophages and FBGCs is localized to the ventral 1 µm of the cells, only up to one third of the F-actin is co-localized with gelsolin. Significantly more co-localization occurred in FBGCs.
To examine the tenacity of podosome-mediated adhesion, FBGC cultures were treated with 5 µM cytochalasin D, which disrupts F-actin. Microfilaments were disrupted throughout the cytoplasm. However, punctate F-actin at the ventral cell surface was resistant to cytochalasin D treatment (Figure 4), suggesting that podosome structural proteins protected the F-actin from depolymerization.
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IL-13-Induced Alterations in Macrophage Morphology Are Culture Surface-dependent
Interestingly, we found that IL-13 induced a dramatic change in macrophage morphology on culture surfaces that did not support macrophage fusion and FBGC formation (Figure 5). When not treated with exogenous cytokines, cultures stained for microtubules and intermediate filaments showed no significant changes in organization throughout the 10-day culture period (not shown). Spread cells contained a centrally located concentration of tubulin from which highly branched and interconnected microtubules radiated towards and parallel to the cell periphery (Figure 5E). An extremely dense intermediate filament network was observed (Figure 5F).
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On coverslips not treated with any silane, IL-13 did not induce FBGC formation, although macrophage adhesion was comparable to that on silane-treated coverslips. Instead, macrophages assumed an elongated, spindled morphology. Where spindles terminated on the coverslip surface, punctate F-actin structures formed (Figure 5A) and were identified as podosomes by vinculin, talin, paxillin, and gelsolin staining (not shown). Microfilaments, microtubules, and intermediate filaments extended along the long axis of these macrophages (Figure 5AC). Microtubules and intermediate filaments were organized into meshworks where spindles terminated on the surface in an organization similar to that of spread macrophages (Figure 5B and Figure 5C compared to Figure 5E and Figure 5F, respectively).
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Discussion |
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The results presented here demonstrate that there is continued cytoskeletal rearrangement during the formation of FBGCs and that there are quantitative differences between the distribution of microfilaments in FBGCs and in macrophages. The increase in F-actin at the ventral cell surface of FBGCs may reflect the more spread morphology compared to macrophages, or it may indicate that FBGCs require more adhesive structures per area for adhesion than macrophages. In support of the latter, the fluorescence staining of punctate F-actin in FBGCs appeared much denser and more intense than that in macrophages. These quantitative differences in microfilament distribution between FBGCs and macrophages, but not overall polymerization as measured by equivalent amounts of total F-actin, suggest that additional functional specialization may be conferred by macrophage fusion to form FBGC. A summary of the results from this study is presented in Table 2.
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The cytoskeletal organization observed in monocytes and macrophages in this study confirms and extends to human monocytes/macrophages earlier reports on nonhuman monocytes/macrophages (
These adhesive structure results are in agreement with previous investigations (
Characterization of adhesive structures may be valuable for interpretation of macrophage function in chronic inflammatory responses. Focal contacts function in the stabilization of cell adhesion to an underlying substrate (
The failure of cytochalasin D to destabilize podosome structure provides insight into the strength of adhesion of spread macrophages and FBGCs to synthetic surfaces. Adhesion strength may result from the combined influences of the multiple protein components of podosomes, which could lend stability to these structures. Macrophage and FBGC trypsin resistance (personal observations) could be similarly explained because the action of trypsin depends on the disruption of F-actin and subsequent rounding of cells (
Our findings also demonstrate that the nature of the culture surface with which monocytes interact substantially influences the subsequent phenotypic response to IL-13. This is similar to results with IL-4 (
Taken together, these studies enhance our understanding of macrophage and FBGC cytoskeletal and adhesive structural support, which is likely critical for the acquisition of functional specializations by these cells during the chronic inflammatory response.
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Acknowledgments |
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Supported by the National Heart, Lung, and Blood Institute, Devices and Technology Branch, Grant HL 55714, the Whitaker Foundation, and the Center for Cardiovascular Biomaterials at Case Western Reserve University.
Received for publication May 20, 1998; accepted September 8, 1998.
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