Phagocytosis stimulates mobilization and ... - Semantic Scholar

Phagocytosis stimulates mobilization and shedding of intracellular CD16A in human monocytes and macrophages: inhibition by HIV-1 infection Nicole L. Webster,*,†,1 Katherine Kedzierska,*,‡ Rula Azzam,*,§ Geza Paukovics,* John Wilson,¶ Suzanne M. Crowe,*,§ and Anthony Jaworowski*,§ *AIDS Pathogenesis Research Program, Macfarlane Burnet Institute for Medical Research and Public Health, Melbourne, Australia; Departments of †Immunology and §Medicine, Monash University, Melbourne, Australia; ‡ Department of Microbiology and Immunology, University of Melbourne, Australia; and ¶Respiratory Unit, The Alfred Hospital, Melbourne, Australia

Abstract: Surface and intracellular staining coupled with flow cytometric analysis was used to show for the first time that human macrophages and a minor subset of peripheral blood monocytes have an internal pool of CD16A, which is mobilized and shed during Fc receptor for immunoglobulin Gmediated phagocytosis. Human immunodeficiency virus type 1 (HIV-1) infection of monocyte-derived macrophages in vitro led to a reduction in the phagocytosis-induced up-regulation in CD16A shedding. These results suggest that monocytes and macrophages may be a source of soluble CD16A, which is elevated in the serum of patients in a variety of disease states and that the mobilization and shedding of CD16A in response to phagocytosis are disrupted by HIV-1 infection. J. Leukoc. Biol. 79: 294 –302; 2006. Key Words: soluble CD16 䡠 Fc␥R 䡠 immunoglobulin G 䡠 soluble Fc␥RIII 䡠 monocyte subsets

INTRODUCTION CD16 [Fc receptor for immunoglobulin G (IgG; Fc␥R)III] is a multifunctional, low/intermediate affinity receptor for the Fc portion of IgG. This receptor is involved in phagocytosis, secretion of enzymes, inflammatory mediators, antibody-dependent cellular cytotoxicity (ADCC), and clearance of immune complexes [1]. CD16 exists in two isoforms encoded by separate homologous genes, which are expressed in a cell typespecific manner [2]. Fc␥RIIIa (CD16A) is encoded by FcRIII-2 and is an intermediate affinity type 1 transmembrane glycoprotein expressed by a subset of T lymphocytes, mast cells, natural killer (NK) cells, monocytes, and macrophages. Fc␥RIIIb (CD16B), encoded by FcRIII-1, is a low-affinity, glycosyl-phosphatidylinositol (GPI)-linked receptor expressed by neutrophils and on eosinophils, in an inducible manner [2]. Studies using knockout mice have shown that the murine homologue of Fc␥RIII is indispensible for macrophage phagocytosis of targets opsonized with IgG1 (but not IgG2a or IgG2b) and for ADDC by NK cells, mast cell degranulation, IgG294

Journal of Leukocyte Biology Volume 79, February 2006

dependent, passive cutaneous anaphylaxis, and Arthus reactions [3]. CD16⫹ monocytes are a minor subset, which typically comprises 5–15% of the peripheral blood monocytes in healthy individuals. The proportion of CD16⫹ monocytes is reportedly expanded in a number of inflammatory conditions including atherosclerosis [4], arthritis [5, 6], sepsis [7, 8], erysipelas [9], pulmonary alveolar proteinosis [10], inflammatory bowel disease [11], as well as in hemodialysis [12], cardiac surgery [13, 14], and with human immunodeficiency virus type-1 (HIV-1) infection [15, 16]. CD16⫹ monocytes in human peripheral blood have been reported to exhibit a mature, macrophage-like morphology [17], to have lower phagocytic and oxidative activity [18] and more efficient antigen presentation [17, 18], and to produce more proinflammatory cytokines {interleukin (IL)-1, IL-6, and tumor necrosis factor-␣ (TNF-␣) [19]} and neurotoxic factors [20] compared with the majority of monocytes, which is phenotypically CD14⫹/CD16 –. As well as being expressed on the surface of cells, CD16 is also found in a soluble form (sCD16), which is detectable in saliva, synovial, seminal fluid, serum, and plasma [21–25] under normal and pathological conditions. The level of sCD16 circulating in plasma of healthy individuals is ⬃1 ␮g/mL [26] and increases in a manner that correlates with disease progression in a number of disease states including rheumatoid arthritis [21, 27, 28], myeloma [29], atherosclerosis [30], and HIV infection [31]. The existence of intracellular pools of CD16B in neutrophils [32–34] and eosinophils [35] and shedding of CD16 by neutrophils [21, 24, 33], eosinophils [35], and NK cells [36, 37] has been well-characterized. However, the contribution of monocytes and macrophages to plasma levels of sCD16 and the presence of an intracellular pool in cells of this lineage require investigation. In this study, we demonstrate the existence of a pool of intracellular CD16A in human monocytes, monocyte-

1

Correspondence: AIDS Pathogenesis and Clinical Research Program, Macfarlane Burnet Institute for Medical Research and Public Health, 85 Commercial Rd., Melbourne 3004, Australia. E-mail: [email protected] Received July 11, 2005; revised September 5, 2005; accepted October 7, 2005; doi: 10.1189/jlb.0705382.

0741-5400/06/0079-294 © Society for Leukocyte Biology

derived macrophages (MDM), and alveolar macrophages and show that this pool is mobilized during Fc␥R-mediated phagocytosis. Furthermore, we show that CD16A appears in the culture medium during Fc␥R-mediated phagocytosis, suggesting that it is secreted as a cleaved product. HIV-1 infection results in reduction of CD16 mobilization and shedding during phagocytosis, revealing a novel mechanism in monocyte and macrophage response to HIV infection.

MATERIALS AND METHODS Isolation and culture of human monocytes Human monocytes were isolated from buffy coats of HIV-, hepatitis B virus -, and human T cell lymphotropic virus-seronegative blood donors (purchased from Red Cross Blood Bank, Melbourne, Australia) by Ficoll-Paque density gradient centrifugation and countercurrent elutriation purification [38] or plastic adherence [39] as described previously. Immediately after isolation, cell viability was greater than 95% as assessed by trypan blue exclusion, and the purity of monocytes was greater than 93% as determined by immunofluorescent staining with anti-CD14 monoclonal antibody (mAb; Becton Dickinson, Mountain View, CA) and flow cytometric analysis. Cells were cultured in Iscove’s modified Dulbecco’s medium (Cytosystem, Castle Hill, Australia) supplemented with 10% heat-inactivated human AB⫹ serum, 2 mM L-glutamine, and 24 ␮g/ml gentamicin (supplemented Iscove’s medium) in suspension in polytetrafluorethylene (Teflon) pots (Savillex, Minnetonka, MN) at an initial concentration of 1 ⫻ 106 cells/ml. Each experiment was performed using MDM from at least three different donors unless otherwise specified.

HIV-1 infection of MDM Infection of MDM with the M-tropic stain HIV-1Ba-L (AIDS Research and Reference Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD) was performed 5 days after monocyte isolation and culture in Teflon pots, as described previously [40]. Phagocytosis assay was performed 7 days postinfection (Day 12 MDM). Controls were mock-infected and cultured under the same conditions. HIV-1 replication was quantified by measuring reverse transcriptase (RT) activity using the micro-RT assay as described previously [40].

Bronchoalveolar lavage Bronchoalveolar lavage (BAL) was performed on healthy individuals with informed consent as described previously [41]. Briefly, a bronchoscope (Olympus, Faulding Imaging, Melbourne, Australia) was placed in wedge position in the middle lobe or lingula. Four 50 ml aliquots of sterile solution were instilled and then aspirated immediately by gentle suction. Samples were pooled in a sterile, siliconized container to prevent adherence of macrophages. Within 3 h of collection, the cells were washed twice in cold (4°C) calcium and magnesium-free, phosphate-buffered saline-cyclophosphamide, methotrexate, and fluorouracil (PBS-CMF) supplemented with 1% fetal calf serum (FCS) at 350 g for 7 min. Supernatant was aspirated to 200 ␮l, and cells were incubated at 4°C for 20 min to reduce autofluorescence. Alveolar macrophages, present in the sample, were then stained for surface and intracellular CD16.

Fc␥R-mediated phagocytosis assays using elutriation-purified monocytes or cultured MDM in suspension Target particles were opsonized immediately prior to the phagocytosis assay as described previously [42]. Briefly, latex beads (3 ␮m in diameter, Sigma Chemical Co., St. Louis, MO) were coated with bovine serum albumin (BSA; Sigma Chemical Co.), washed, and opsonized with rabbit anti-BSA antiserum (ICN-Cappel, Aurora, OH). Elutriation-purified monocytes or MDM cultured for 7 days post-isolation were dispensed into polypropylene tubes (Becton Dickinson) at 2 ⫻ 106, washed twice in PBS-CMF (500 g, 5 min), and cooled on ice in 100 ␮l PBS-CMF for 20 min. To perform Fc␥R-mediated phagocy-

tosis, MDM were incubated with or without opsonized beads at 37°C at varying cell-to-particle ratios (1:5 to 1:50) in duplicate. At various time-points, phagocytosis was arrested by plunging the tubes into ice. Cells were pelleted, supernatant stored, and cells fixed with 1 ml of 3% ultrapure formaldehyde (Polysciences, Warrington, PA) in PBS-CMF for 10 min at 4°C, followed by two washes with cold 0.1 M glycine in PBS-CMF and staining for surface and intracellular CD16.

Expression of surface and intracellular CD16 by purified cell populations Purified monocytes, cultured MDM, and alveolar macrophages (or monocytes and MDM fixed following phagocytosis as described above) were stained for surface and intracellular CD16 as described previously [35]. First, cells were stained for expression of surface CD16 with anti-CD16 mAb conjugated to fluorescein isothiocyanate (FITC; Serotec, Raleigh, NC) or isotype-matched control IgG1 conjugated to FITC (Becton Dickinson) as well as for the specific cell markers CD14 (for monocytes/macrophages), CD56 (for NK cells), or CD3 (for T lymphocytes; Becton Dickinson), each conjugated to phycoerythrin (PE). After 30 min incubation at 4°C, cells were washed with cold PBS-CMF (500 g, 5 min, 4°C), fixed with 1 ml 3% formaldehyde in PBS-CMF for 10 min at 4°C, washed twice with cold 0.1 M glycine in PBS-CMF, and permeabilized with 0.1% Triton X-100 (Merck, Kilsyth, Australia) for 1 min. After two washes with 5% FCS in PBS, cell nonspecific-binding sites were blocked with 10% mouse serum for 20 min at 4°C and washed twice with 1% FCS in PBS-CMF. Cells were stained for intracellular CD16 levels with anti-CD16 mAb conjugated to Cy-Chrome or isotype-matched control IgG1 conjugated to Cy-Chrome (BD PharMingen, San Diego, CA) for 30 min on ice, followed by a wash with cold 1% FCS/PBS-CMF. Cells were fixed with 200 ␮l 1% formaldehyde, and the expression of surface and intracellular CD16 in phagocytosing or resting cells was analyzed by flow cytometry (FACStarPlus, Becton Dickinson). Similar antibody staining was obtained when MDM were fixed with formaldehyde prior to or after the addition of antibody. The CD16A mAb used in this study were generated from the same clone (3G8) and displayed nonblocking characteristics.

Expression of surface and intracellular CD16 in whole blood by flow cytometry Blood (5 mL) was collected into EDTA vacutainers, 250 ␮L blood was added into polypropylene tubes (Becton Dickinson) and washed with PBS-CMF (500 g, 5 min), and red blood cells were lysed with 0.5 mL ammonium chloride/ EDTA lysis buffer (150 mM NH4Cl, 10 mM NaHCO3, and 1 mM EDTA) for 5 min at 37°C. Leukocytes were washed twice with PBS-CMF (500 g, 5 min) and stained for surface and intracellular CD16 as described above. Forward-scatter and side-scatter (FSC and SSC) profiles were used to define monocyte and lymphocyte populations. Within the monocyte population gate, CD14⫹ cells were identified, and surface and intracellular CD16 expression levels were determined. In the lymphocyte gate, CD3⫹ cells and CD56⫹ cells were identified, and their surface and intracellular CD16 expression levels were determined.

Analysis of CD16A secretion during Fc␥R-mediated phagocytosis Culture supernatants were collected immediately following phagocytosis assays using elutriation-purified monocytes or cultured MDM, performed as described above. Cell lysates were prepared using 1% Triton X-100 in 25 mM Tris, pH 7.5, buffer supplemented with the following protease inhibitors: 1 mM pefabloc, 1 ␮M pepstatin, 1 ␮M leupeptin (Boehringer-Mannheim, Mannheim, Germany). Cell extracts containing 30 ␮g protein and 20 ␮L culture medium were separated on 10% sodium dodecyl sulfate (SDS)-polyacrylamide gels under nonreducing conditions. Protein was transferred to nitrocellulose membrane, and immunoblotting was performed. Membranes were blocked for 1 h at room temperature with Odyssey blocking buffer (927-40000, LiCOR Biosciences, Lincoln, NE) and incubated with mouse anti-CD16A mAb (HuNK2, Neomarkers, Union City, CA) overnight at 4°C. Following washing with 0.1% Tween 20 in PBS, antibody binding was detected using the secondary goat anti-mouse mAb conjugated to Alexa Fluor 680 (A-21057, Molecular Probes, Eugene, OR). Bands were detected using the LiCOR Odyssey infrared imaging system. Alternatively, a goat anti-mouse mAb secondary conjugated to horse-

Webster et al. CD16 shedding by monocytes and macrophages

295

radish peroxidase (HRP) was used to detect anti-CD16 mAb binding, and HRP substrate was developed by enhanced chemiluminescence (ECL) where indicated.

RESULTS Human monocytes and macrophages contain an internal pool of CD16A The expression of intracellular CD16A by monocytes (CD14⫹), T lymphocytes (CD3⫹), and NK cells (CD56⫹) was examined by three-color immunofluorescence analysis of whole blood. Cell populations were identified by the relevant PE-labeled mAb. The staining strategy described in the Materials and Methods is specific for differentially detecting surface CD16 compared with intracellular CD16 on the same cell, using the mAb FITC-labeled anti-CD16 and Cy-Chrome-labeled anti-CD16, respectively. Both of these anti-CD16 mAb are clone 3G8. Control experiments showed that in the absence of cell permeabilization, the surface staining of cells with FITC-labeled anti-CD16 was saturating, as there was no binding of the Cy-Chrome-labeled anti-CD16 mAb above background levels as indicated by matched isotype control. Reversing the order of staining, using CD16-CyC for surface CD16 staining and CD16-FITC for intracellular CD16 staining gave similar results. Thus, this staining combination differentiated surface and intracellular expression of CD16. Using this staining protocol, a subset of monocytes was found to contain an intracellular pool of CD16 (Fig. 1a). Of the minor proportion of the CD14⫹ monocytes, which express CD16 on their surface (24.3⫾4.9, n⫽3), 35% of these cells also contained an intracellular pool of CD16 (8.5⫾3.6, n⫽3). As it is known that monocytes and macrophages specifically express CD16A, this isoform will be referred to in the following text, although the antibody used here does not discriminate between CD16A and CD16B. The monocytes containing intracellular CD16A were all positive for CD16A surface expression and also expressed the highest levels of surface CD16A (Fig. 1b). None of the CD14⫹ monocytes, which lack CD16A surface expression, were found to have an intracellular pool of CD16A (Fig. 1b). The size and granularity of monocytes expressing surface and intracellular CD16A were greater than that of the monocyte population expressing only surface CD16A and of the major CD14⫹ monocyte population, which lacks CD16A expression (Fig. 1c). Whereas only a minor subset of freshly isolated monocytes expressed intracellular CD16A, mature MDM expressed high levels of intracellular CD16A when cultured for 7 days (mean: 74.1%, range: 51.4 – 88.14%, n⫽5), indicating the presence of a significant intracellular pool of CD16A in MDM. The level of intracellular CD16A in MDM was examined at various times after monocyte culture (Fig. 2a). Intracellular CD16A was low in freshly isolated monocytes and MDM cultured for 2 days and was increased progressively between Days 2 and 7 of culture. A similar expression pattern for CD16A was observed in alveolar macrophages, which were assessed ex vivo within 3 h of the lavage procedure. Human tissue macrophages recovered 296


from BAL displayed high levels of surface and intracellular CD16A (75.0% and 79.3%, respectively; Fig. 2b). To further support the existence of an intracellular pool of CD16A, as has been shown with CD16B in neutrophils [33] and eosinophils [35], NK and T cells in whole blood were also analyzed, and subsets were found to contain intracellular CD16. The majority of CD56⫹ NK cells expresses CD16A on their surface (83.5⫾12.8, n⫽3), and 47% of these cells also express intracellular CD16A (39.5⫾4.6, n⫽3; Fig. 1a). Similarly, we also found that a minor subset of CD3⫹ T lymphocytes expresses surface CD16A (24.2⫾14.9, n⫽3), and 65% of this subset contains detectable, intracellular CD16A (15.7⫾5.5, n⫽3). These results show that detectable, intracellular CD16A pools exist in subsets of NK cells, T lymphocytes, and monocytes as well as most macrophages.

Internal CD16A is mobilized during Fc␥R-mediated phagocytosis As the main function of CD16A on monocytes and macrophages is to recognize IgG-opsonized targets, we investigated the fate of surface and intracellular CD16A in human MDM during Fc␥R phagocytosis. MDM cultured for 7 days were incubated with latex beads coated with BSA and opsonized with rabbit anti-BSA antiserum. Addition of IgG-opsonized targets resulted in a progressive decline in levels of intracellular CD16A in a time-dependent manner, which was proportional to the ratio of phagocytic target particles to MDM (Fig. 3a). At a ratio of 5:1, the internal pool of CD16A was depleted within 30 min, whereas at a ratio of 50:1, intracellular CD16A was undetectable within 5 min (Fig. 3a). Simultaneous measurement of cell-surface CD16A failed to detect a concomitant increase in expression during Fc␥R-mediated phagocytosis.

Appearance of sCD16A during Fc␥R-mediated phagocytosis As mobilization of intracellular CD16A did not result in a measurable increase in surface CD16A on MDM, the possibility that phagocytosis resulted in shedding of the receptor was investigated. Peripheral blood monocytes purified by elutriation on the day of venepuncture (d0) and cultured MDM (d5) were incubated with and without IgG-opsonized latex beads for 30 min at 37°C. Culture supernatant was collected, and samples were analyzed by immunoblotting for CD16A. A broad band of molecular weight ⬃60 kD was detected in the medium 30 min after exposure of MDM to latex beads, coincident with a decrease in intracellular CD16A, suggesting that CD16A is secreted as a cleaved product (Fig. 4a). There was little CD16A detected in medium following incubation in the absence of latex beads, suggesting a low rate of basal shedding (Fig. 4b). CD16A was detectable in the culture supernatant as early as 2 min following exposure to the latex bead targets (Fig. 4a). No CD16A was detected in any of the culture supernatants when cells were incubated at 4°C, with or without the latex beads (data not shown). Shedding of CD16A by freshly isolated human monocytes was also demonstrated using the sensitive infrared detection methods (Fig. 4c). http://www.jleukbio.org

Fig. 1. Flow cytometric analysis of surface and intracellular CD16A expression in human monocytes, T cells, and NK cells. Whole blood (250 ␮L) was stained for CD14, CD56, and CD3 expression to identify monocytes, NK cells, and T cells, respectively. Cell populations were defined by first gating on the monocyte or lymphocyte population based on the FSC versus SSC profile and then specific CD14, CD3, or CD56 marker-positive cells. (a) Surface CD16A expression was quantified using anti-CD16 mAb conjugated to FITC (shaded bars), and intracellular CD16A was quantified using anti-CD16 mAb conjugated to Cy-Chrome (solid bars). The results are expressed as percentage of CD16A-positive cells within the specific cell population and are corrected for background fluorescence (matched isotype control staining). Data shown represent means ⫾ SD from three experiments. (b) Quadrant graph showing a representative example of the expression of surface and intracellular CD16A expression by CD14⫹ monocytes in whole blood. Lower left quadrant shows CD16A surface and intracellular negative monocytes (⫺/⫺); the lower right quadrant, surface-positive and intracellular CD16A-negative monocytes (⫹/⫺); and the upper right quadrant, monocytes expressing surface and intracellular CD16A (⫹/⫹). No monocytes express intracellular CD16A in the absence of surface CD16A (upper left quadrant). The same results were observed when the staining order was reversed, and surface CD16A was detected with CyC-conjugated anti-CD16 mAb, and intracellular CD16 was detected with FITC-conjugated anti-CD16 mAb. (c) Size and granularity distribution of the monocyte populations. The solid black histogram represents the surface and intracellular CD16A-positive monocytes (⫹/⫹), the black line shows the surface CD16A-positive and intracellular-negative monocytes (⫹/⫺), and the dotted line shows the CD16A surface and intracellular-negative monocytes (⫺/⫺).

HIV-1 infection of human MDM inhibits phagocytosis-stimulated CD16A shedding The above data suggest that mobilization and shedding of CD16A are integral parts of the phagocytic process. As we have shown that phagocytosis by monocytes and macrophages is inhibited by HIV-1, whereas CD16 expression is not affected, we examined the impact of HIV-1 infection of MDM on CD16A shedding. MDM were infected or mock-infected after 5 days in culture with the M-tropic strain HIV-1Ba-L at a multiplicity of infection of 0.1–1.0, which we have shown inhibits Fc␥R-mediated phagocytosis by ⬃50% [43]. After 7 days infection, MDM were incubated with IgG-opsonized latex

beads as a phagocytic target for 30 min at a target-to-MDM ratio of 10:1, and then culture medium was collected and analyzed by immunoblotting for sCD16A. The level of total CD16A expression (surface and intracellular) was investigated on these cells following staining of permeabilized cells and flow cytometric analysis. HIV-1 infection of the MDM was confirmed by RT analysis of culture supernatants taken immediately prior to phagocytosis. The increased shedding of CD16A compared with the notarget control was compared between HIV-1-infected and uninfected MDM (Fig. 5, a and b). HIV-1 infection resulted in a threefold reduction in the increase in shedding of CD16A


297

Fig. 2. An intracellular pool of CD16A is present in human monocytes and macrophages. (a) Altered expression of intracellular CD16A accompanies differentiation of monocytes into monocyte-derived macrophages. Monocytes isolated from buffy coats were cultured for up to 7 days in Teflon pots. Intracellular CD16A was assessed by flow cytometry following culture for 2, 4, and 7 days. Monocytes/MDM were stained with anti-CD16 conjugated to FITC (surface), fixed, permeabilized, and stained with anti-CD16 conjugated to Cy-Chrome (intracellular) mAb as described in Materials and Methods. A representative result (from n⫽3 donors) is shown of intracellular staining for CD16 with CD16-CyC (solid line histograms) or isotypematched control (dotted line histograms) with net mean fluorescence intensity (nMFI; corrected for background fluorescence as determined by isotypematched control) stated. Dot quadrant plots show surface and intracellular expression of CD16. (b) Surface and intracellular expression of CD16A by bronchoalveolar macrophages ex vivo. Freshly isolated bronchoalveolar macrophages were stained for surface and intracellular expression of CD16A as in a. Cells were labeled with isotype-matched control mAb (dotted line histograms) or anti-CD16 mAb conjugated to FITC for the surface staining and anti-CD16 mAb conjugated to Cy-Chrome for the intracellular staining (black histograms). Dot quadrant plots show surface and intracellular expression of CD16 by the bronchoalveolar macrophages.

during phagocytosis (P⫽0.05, n⫽3). No significant difference was observed in the basal level of shedding of CD16A between uninfected and HIV-infected MDM, showing that inhibition is specific for events downstream of phagocytosis. It has been shown previously that surface CD16 expression is not altered on Day 12 MDM by HIV infection (7 days post-infection) [43]. A 60% reduction in the total level of CD16 expression was observed in the HIV-infected MDM compared with the uninfected MDM (Fig. 5c), indicating a correlation between the available pool of CD16 and the level of CD16 shedding during phagocytosis.

DISCUSSION In this study, we have shown that monocytes and macrophages have an intracellular pool of CD16A, which is mobilized and 298


shed during phagocytosis. In addition, we have demonstrated that HIV infection leads to a decrease in CD16A mobilization and shedding during phagocytosis. Only a minor subset of surface CD16-expressing monocytes also contains an intracellular pool of CD16. These cells, with an intracellular pool of CD16, are larger and more granular than both of the CD16⫹ monocytes, which lack detectable levels of intracellular CD16 and the major CD14⫹/CD16 – monocyte population. Previous investigators using elutriated monocytes reported that monocytes expressing CD16 on their surface are smaller and less granular than the CD16 – monocytes [18, 44]. The anticoagulant used and subsequent processing of the monocytes can affect CD16 expression. To avoid artifacts of monocyte isolation, we performed phenotypic analysis of monocytes present in freshly drawn whole blood. We observed heterogeneity in the size of surface CD16-expressing http://www.jleukbio.org

Fig. 3. Disappearance of intracellular CD16 in MDM during phagocytosis is not associated with an increase in surface CD16 expression. Phagocytosis was performed using MDM on Day 7 post-isolation. MDM were incubated with or without opsonized latex beads at 37°C at varying cell-to-bead ratios (from five latex beads per MDM to 50 latex beads per MDM) in duplicate. At 2, 5, 15, or 30 min, phagocytosis was arrested by plunging the tubes into ice. Cells were fixed with 1 ml 3% ultrapure formaldehyde and staining for surface and intracellular CD16 using anti-CD16 mAb. These data represent means ⫾ SD. Results are representative of three experiments using MDM from different donors.

monocytes, ranging from the smaller, less granular monocytes through to larger and more granular monocytes. Differences in detection of CD16 expression in these studies highlight the dynamic nature of expression of the CD16 receptor. It is notable that the major CD14⫹ monocyte population lacks an intracellular CD16 pool (Fig. 1b) as detected by flow cytometry. Mobilization and shedding of the intracellular CD16A pool in monocytes and macrophages are caused by a variety of stimuli. We have demonstrated a novel role for phagocytosis in this process. Little is known about shedding of CD16 from monocytes and macrophages. Levy and colleges [45] described shedding of CD16 by alveolar macrophages during in vitro culture and showed that the matrix metalloproteinase (MMP) inhibitor 1,10-phenanthroline, could inhibit 70% of the observed shedding. When the same inhibitor was tested in the current study, it was also unable to abrogate all shedding by monocytes during phagocytosis. CD16A was detected by immunoblotting in the culture supernatant of monocytes, which

had phagocytosed latex bead targets, even in the presence of the MMP inhibitor (data not shown). Taken together, these observations suggest that MMPs are only one mechanism involved in proteolysis of CD16 by monocytes and macrophages. Other mechanisms may include microvesicular shedding as observed with exosome vesicular shedding of soluble cytokine receptors [46] or shedding following proteolysis by other proteinases (reviewed in ref. [47]). The results presented above show that surface expression of CD16A on monocytes is highly dynamic. It has been well established that cytokines such as IL-1 and TNF-␣ increase surface expression of CD16A on monocytes [48, 49], whereas transforming growth factor-␤ results in a decrease in CD16A expression [50 –52]. An in vitro transendothelial migration model, which uses human umbilical vein endothelial cell monolayer on collagen [53], has demonstrated two-thirds of the CD16⫹ monocytes can migrate into tissue within 2 h, and these cells down-regulate expression of CD16. Fifty percent of these monocytes reverse-transmigrate out of the collagen after 2 days, up-regulating surface CD16 expression upon leaving the tissue. Migration of monocytes into tissues and migration in the presence of phagocytic target particles result in rapid down-regulation of surface CD16A on monocytes [53]. Our results are constistent with this and suggest that this downregulation of receptor expression is more likely to be a result of shedding of CD16A from the surface rather than internalization. The existence of an intracellular pool of CD16B and its shedding following cell activation and during apoptosis have been well-characterized in neutrophils. Stimulation of neutrophils, with formyl-Met-Leu-Phe or phorbol 12-myristate 13acetate (PMA) [33, 54], causes rapid translocation of intracellular CD16B from its storage compartments as well as surface shedding of the receptor. This increased surface expression is only transiently observed [33]. The CD16B cleavage can be inhibited by serine protease inhibitors, which target the proteases present within the azurophilic granules, and MMP inhibitors, depending on the stimuli used to activate the cell [54, 55]. The cleavage site was determined to be between Val 196 and Ser 197 at the C-teminal sequence, close to the membrane GPI anchor [24]. The nature of the compartment containing the newly discovered pool of CD16A in monocytes and macrophages and the mechanism of its mobilization and shedding require characterization. The majority of sCD16 in plasma is believed to be derived from neutrophils [56, 57] and to a lesser extent, NK cells [21], although the contribution of monocytes and macrophages to the sCD16 in plasma has not been determined. In recent studies using a newly developed, macrophage-specific anti-CD16A mAb, it has been shown that CD16A is shed from monocytes and macrophages in plasma of people with inflammatory conditions such as rheumatoid arthritis [27] and atherosclerosis [30]. Our data support the view that monocytes and macrophages contribute to the plasma levels of sCD16. CD16⫹ monocytes may play an important role in HIV pathogenesis. An increase in the proportion of monocytes with this phenotype has been described in the peripheral blood of HIV-infected patients with HIV-associated dementia (HAD) [20]. CD16⫹ monocytes also accumulate in perivascular sites


299

Fig. 4. Secretion of CD16A by monocytes and MDM during Fc␥R-mediated phagocytosis. (a) Day 7 MDM (2⫻106) were incubated in polypropylene tubes at 37°C in PBS-CMF with IgG-opsonized 3 ␮m latex beads, at a bead-to-cell ratio of 10:1 for 0, 2, or 30 min. Phagocytosis was stopped by plunging tubes on ice, and the culture medium was collected and clarified by centrifugation (20,000 g, 10 min). Samples of cell extracts containing 30 ␮g protein and 20 ␮l clarified culture medium were analyzed by SDS-polyacrylamide gel electrophoresis (PAGE). Samples of each culture medium (20 ␮l) were resolved by SDS-PAGE under nonreducing conditions, probed with a mouse anti-CD16 mAb (HuNK2, Neomarkers) overnight, followed by a secondary anti-mouse mAb conjugated to HRP and ECL detection. MDM lysates represent total CD16 in the extractable cellular protein, and supernatant represents CD16 shed into the culture media. (b) MDM were incubated with beads as in panel (a) for 30 min with or without the opsonized latex bead phagocytic target. Clarified culture medium (20 ␮l) was analyzed by SDS-PAGE. The immunoblot shown is representative of three experiments using MDM prepared from different donors. (c) Sensitive LiCOR detection of CD16 shed into culture supernatant by monocytes. Elutriation-purified monocytes (2⫻106) were incubated at 37°C at 100 ␮l/tube in PBS-CMF ⫾ IgG-opsonized latex beads at a bead-to-cell ratio of 10:1 for 30 min. Culture medium (20 ␮l) was analyzed as in panel (a) except that a secondary anti-mouse mAb conjugated to Alexa Fluor 680 was used, and bands were detected using the LiCOR Odyssey infrared imaging system.

in the brain of patients with HAD [58]. Monocytes from HIVinfected individuals harbor replication-competent virus [59], and the CD16⫹ monocyte subset has an increased susceptibility to infection in vitro as well as in vivo [60]. The CD16⫹

monocytes may contribute to maintaining tissue macrophage reservoirs of HIV [61, 62]. Current, highly active, antiretroviral therapy is unable to eradicate infection, and sanctuary sites such as the brain are particularly resistant.

Fig. 5. CD16A shedding induced during phagocytosis by MDM is reduced significantly in HIV-1-infected MDM. Day 12 MDM (uninfected or 7 days post-HIV-1Ba-L infection) were incubated at 37°C in 400 ␮l PBS-CMF ⫾ IgG-opsonixed latex beads (3 ␮m diameter) at a bead-to-cell ratio of 10 latex beads per MDM for 30 min. Phagocytosis was stopped by placing cultures on ice, and culture medium was collected and clarified. Culture medium was resolved by SDS-PAGE under nonreducing conditions, probed with a mouse anti-CD16 mAb (HuNK2, Neomarkers) overnight, followed by a secondary anti-mouse mAb conjugated to Alexa Fluor 680. (a) Band intensities on the Western blots were analyzed using the LiCOR Odyssey infrared imaging system analysis software with lane backgrounds subtracted. A representative Western blot is shown (b) The graph represents the results (mean⫾SE) of the percentage increase in CD16A shedding during phagocytosis of latex beads compared with the basal level of shedding at 37°C (no bead control), n ⫽ 3. Significance was determined using paired Student’s t-test. (c) Reduction in total CD16 expression levels was observed by the HIV-infected MDM compared with the uninfected MDM at Day 12. Prior to the phagocytosis, an aliquot of uninfected and HIV-infected MDM was fixed, permeablized, and stained for CD16 expression. The total CD16 expression (surface and intracellular) by the uninfected CD16 was normalized to 100% for each donor, and the level of CD16 expression by the HIV-infected MDM was compared with the matched donor’s uninfected control.

300


http://www.jleukbio.org

Susceptibility of HIV-infected individuals to opportunistic infections is largely a result of impaired function of cells of the macrophage lineage, as these cells control the majority of opportunistic pathogens via phagocytosis and intracellular killing. We have shown that HIV-1 infection interferes with the process of CD16 mobilization and shedding during phagocytosis. Impaired CD16 mobilization may contribute to defective phagocytosis of IgG-opsonized, opportunistic pathogens [43], and limiting shedding of sCD16 would potentially contribute to inflammation in the brain, given the known function of soluble, low-affinity IgG receptors in inhibiting immune complex-mediated inflammation [63, 64]. An increase in sCD16 can be detected in plasma early in HIV-1 infection followed by a decline in levels with progression toward AIDS [31]. Taken together with the results of this study, we propose that progression toward AIDS results in reduction of intracellular pools of CD16 in monocytes and macrophages, contributing to defective phagocytosis of opportunistic pathogens and reduction in phagocytosis-induced up-regulation of shedding of CD16. This may provide a mechanism for the decrease in CD16 levels in the plasma observed with progression toward AIDS.

ACKNOWLEDGMENTS This work was supported in part by funding from the National Health and Medical Research Council (NHMRC) and by the Macfarlane Burnet Centre Research Fund. K. K. was a recipient of a NHMRC Peter Doherty Postdoctoral Fellowship and R. A., an Australian Postgraduate Award. S. M. C. is a NHMRC Principal Research Fellow. N. L. W. and K. K. contributed equally.

REFERENCES 1. van de Winkel, J. G., Anderson, C. L. (1991) Biology of human immunoglobulin G Fc receptors. J. Leukoc. Biol. 49, 511–524. 2. Edberg, J. C., Kimberly, R. P. (1997) Cell type-specific glycoforms of Fc ␥ RIIIa (CD16): differential ligand binding. J. Immunol. 159, 3849 – 3857. 3. Hazenbos, W. L., Gessner, J. E., Hofhuis, F. M., Kuipers, H., Meyer, D., Heijnen, I. A., Schmidt, R. E., Sandor, M., Capel, P. J., Daeron, M., van de Winkel, J. G., Verbeek, J. S. (1996) Impaired IgG-dependent anaphylaxis and Arthus reaction in Fc ␥ RIII (CD16) deficient mice. Immunity 5, 181–188. 4. Draude, G., von Hundelshausen, P., Frankenberger, M., Ziegler-Heitbrock, H. W., Weber, C. (1999) Distinct scavenger receptor expression and function in the human CD14(⫹)/CD16(⫹) monocyte subset. Am. J. Physiol. 276, H1144 –H1149. 5. Kawanaka, N., Yamamura, M., Aita, T., Morita, Y., Okamoto, A., Kawashima, M., Iwahashi, M., Ueno, A., Ohmoto, Y., Makino, H. (2002) CD14⫹,CD16⫹ blood monocytes and joint inflammation in rheumatoid arthritis. Arthritis Rheum. 46, 2578 –2586. 6. Iwahashi, M., Yamamura, M., Aita, T., Okamoto, A., Ueno, A., Ogawa, N., Akashi, S., Miyake, K., Godowski, P. J., Makino, H. (2004) Expression of Toll-like receptor 2 on CD16⫹ blood monocytes and synovial tissue macrophages in rheumatoid arthritis. Arthritis Rheum. 50, 1457–1467. 7. Fingerle, G., Pforte, A., Passlick, B., Blumenstein, M., Strobel, M., Ziegler-Heitbrock, H. W. (1993) The novel subset of CD14⫹/CD16⫹ blood monocytes is expanded in sepsis patients. Blood 82, 3170 –3176. 8. Blumenstein, M., Boekstegers, P., Fraunberger, P., Andreesen, R., Ziegler-Heitbrock, H. W., Fingerle-Rowson, G. (1997) Cytokine production precedes the expansion of CD14⫹CD16⫹ monocytes in human sepsis: a case report of a patient with self-induced septicemia. Shock 8, 73–75.

9. Horelt, A., Belge, K. U., Steppich, B., Prinz, J., Ziegler-Heitbrock, L. (2002) The CD14⫹CD16⫹ monocytes in erysipelas are expanded and show reduced cytokine production. Eur. J. Immunol. 32, 1319 –1327. 10. Yoshioka, Y., Ohwada, A., Harada, N., Satoh, N., Sakuraba, S., Dambara, T., Fukuchi, Y. (2002) Increased circulating CD16⫹ CD14dim monocytes in a patient with pulmonary alveolar proteinosis. Respirology 7, 273–279. 11. Weber, C., Belge, K. U., von Hundelshausen, P., Draude, G., Steppich, B., Mack, M., Frankenberger, M., Weber, K. S., Ziegler-Heitbrock, H. W. (2000) Differential chemokine receptor expression and function in human monocyte subpopulations. J. Leukoc. Biol. 67, 699 –704. 12. Nockher, W. A., Scherberich, J. E. (1998) Expanded CD14⫹ CD16⫹ monocyte subpopulation in patients with acute and chronic infections undergoing hemodialysis. Infect. Immun. 66, 2782–2790. 13. Fingerle-Rowson, G., Auers, J., Kreuzer, E., Fraunberger, P., Blumenstein, M., Ziegler-Heitbrock, L. H. (1998) Expansion of CD14⫹CD16⫹ monocytes in critically ill cardiac surgery patients. Inflammation 22, 367–379. 14. Stefanou, D. C., Asimakopoulos, G., Yagnik, D. R., Haskard, D. O., Anderson, J. R., Philippidis, P., Landis, R. C., Taylor, K. M. (2004) Monocyte Fc ␥ receptor expression in patients undergoing coronary artery bypass grafting. Ann. Thorac. Surg. 77, 951–955. 15. Thieblemont, N., Weiss, L., Sadeghi, H. M., Estcourt, C., HaeffnerCavaillon, N. (1995) CD14lowCD16high: a cytokine-producing monocyte subset which expands during human immunodeficiency virus infection. Eur. J. Immunol. 25, 3418 –3424. 16. Amirayan-Chevillard, N., Tissot-Dupont, H., Capo, C., Brunet, C., DignatGeorge, F., Obadia, Y., Gallais, H., Mege, J. L. (2000) Impact of highly active anti-retroviral therapy (HAART) on cytokine production and monocyte subsets in HIV-infected patients. Clin. Exp. Immunol. 120, 107– 112. 17. Ziegler-Heitbrock, H. W., Fingerle, G., Strobel, M., Schraut, W., Stelter, F., Schutt, C., Passlick, B., Pforte, A. (1993) The novel subset of CD14⫹/ CD16⫹ blood monocytes exhibits features of tissue macrophages. Eur. J. Immunol. 23, 2053–2058. 18. Passlick, B., Flieger, D., Ziegler-Heitbrock, H. W. (1989) Identification and characterization of a novel monocyte subpopulation in human peripheral blood. Blood 74, 2527–2534. 19. Frankenberger, M., Sternsdorf, T., Pechumer, H., Pforte, A., ZieglerHeitbrock, H. W. (1996) Differential cytokine expression in human blood monocyte subpopulations: a polymerase chain reaction analysis. Blood 87, 373–377. 20. Pulliam, L., Gascon, R., Stubblebine, M., McGuire, D., McGrath, M. S. (1997) Unique monocyte subset in patients with AIDS dementia. Lancet 349, 692– 695. 21. de Haas, M., Kleijer, M., Minchinton, R.M., Roos, D., von dem Borne, A. E. (1994) Soluble Fc ␥ RIIIa is present in plasma and is derived from natural killer cells. J. Immunol. 152, 900 –907. 22. Huizinga, T. W., de Haas, M., van Oers, M. H., Kleijer, M., Vile, H., van der Wouw, P. A., Moulijn, A., van Weezel, H., Roos, D., von dem Borne, A. E. (1994) The plasma concentration of soluble Fc-␥ RIII is related to production of neutrophils. Br. J. Haematol. 87, 459 – 463. 23. Sautes, C., Teillaud, C., Mazieres, N., Tartour, E., Bouchard, C., Galinha, A., Jourde, M., Spagnoli, R., Fridman, W. H. (1992) Soluble Fc ␥ R (sFc ␥ R): detection in biological fluids and production of a murine recombinant sFc ␥ R biologically active in vitro and in vivo. Immunobiology 185, 207–221. 24. Galon, J., Moldovan, I., Galinha, A., Provost-Marloie, M. A., Kaudewitz, H., Roman-Roman, S., Fridman, W. H., Sautes, C. (1998) Identification of the cleavage site involved in production of plasma soluble Fc ␥ receptor type III (CD16). Eur. J. Immunol. 28, 2101–2107. 25. Koene, H. R., de Haas, M., Kleijer, M., Roos, D., von dem Borne, A. E. (1996) NA-phenotype-dependent differences in neutrophil Fc ␥ RIIIb expression cause differences in plasma levels of soluble Fc ␥ RIII. Br. J. Haematol. 93, 235–241. 26. Teillaud, J. L., Bouchard, C., Astier, A., Teillaud, C., Tartour, E., Michon, J., Galinha, A., Moncuit, J., Mazieres, N., Spagnoli, R., et al. (1994) Natural and recombinant soluble low-affinity Fc ␥ R: detection, purification, and functional activities. Immunomethods 4, 48 – 64. 27. Masuda, M., Morimoto, T., Kobatake, S., Nishimura, N., Nakamoto, K., Dong, X. H., Komiyama, Y., Ogawa, R., Takahashi, H. (2003) Measurement of soluble Fc␥ receptor type IIIa derived from macrophages in plasma: increase in patients with rheumatoid arthritis. Clin. Exp. Immunol. 132, 477– 484. 28. Masuda, M., Morimoto, T., De Haas, M., Nishimura, N., Nakamoto, K., Okuda, K., Komiyama, Y., Ogawa, R., Takahashi, H. (2003) Increase of soluble Fc␥RIIIa derived from natural killer cells and macrophages in plasma from patients with rheumatoid arthritis. J. Rheumatol. 30, 1911– 1917.


301

29. Mathiot, C., Mary, J. Y., Tartour, E., Facon, T., Monconduit, M., Grosbois, B., Pollet, J. P., Michaux, J. L., Euller Ziegler, L., Sautes, C., Bataille, R., Fridman, W. H. (1996) Soluble CD16 (sCD16), a marker of malignancy in individuals with monoclonal gammopathy of undetermined significance (MGUS). Br. J. Haematol. 95, 660 – 665. 30. Masuda, M., Takahashi, H. (2003) [Soluble Fc ␥ RIIIa derived from macrophages.] Rinsho Byori 51, 1102–1105. 31. Khayat, D., Soubrane, C., Andrieu, J. M., Visonneau, S., Eme, D., Tourani, J. M., Beldjord, K., Weil, M., Fernandez, E., Jacquillat, C. (1990) Changes of soluble CD16 levels in serum of HIV-infected patients: correlation with clinical and biologic prognostic factors. J. Infect. Dis. 161, 430 – 435. 32. Borregaard, N., Miller, L. J., Springer, T. A. (1987) Chemoattractantregulated mobilization of a novel intracellular compartment in human neutrophils. Science 237, 1204 –1206. 33. Tosi, M. F., Zakem, H. (1992) Surface expression of Fc ␥ receptor III (CD16) on chemoattractant-stimulated neutrophils is determined by both surface shedding and translocation from intracellular storage compartments. J. Clin. Invest. 90, 462– 470. 34. Detmers, P. A., Zhou, D., Powell, D., Lichenstein, H., Kelley, M., Pironkova, R. (1995) Endotoxin receptors (CD14) are found with CD16 (Fc ␥ RIII) in an intracellular compartment of neutrophils that contains alkaline phosphatase. J. Immunol. 155, 2085–2095. 35. Zhu, X., Hamann, K. J., Munoz, N. M., Rubio, N., Mayer, D., Hernrreiter, A., Leff, A. R. (1998) Intracellular expression of Fc ␥ RIII (CD16) and its mobilization by chemoattractants in human eosinophils. J. Immunol. 161, 2574 –2579. 36. Harrison, D., Phillips, J. H., Lanier, L. L. (1991) Involvement of a metalloprotease in spontaneous and phorbol ester-induced release of natural killer cell-associated Fc ␥ RIII (CD16-II). J. Immunol. 147, 3459 –3465. 37. Mota, G., Moldovan, I., Calugaru, A., Hirt, M., Kozma, E., Galatiuc, C., Brasoveanu, L., Boltz-Nitulescu, G. (2004) Interaction of human immunoglobulin G with CD16 on natural killer cells: ligand clearance, Fc␥RIIIA turnover and effects of metalloproteinases on Fc␥RIIIA-mediated binding, signal transduction and killing. Scand. J. Immunol. 59, 278 –284. 38. Wahl, L. M., Katona, I. M., Wilder, R. L., Winter, C. C., Haraoui, B., Scher, I., Wahl, S. M. (1984) Isolation of human mononuclear cell subsets by counterflow centrifugal elutriation (CCE). I. Characterization of Blymphocyte-, T-lymphocyte-, and monocyte-enriched fractions by flow cytometric analysis. Cell. Immunol. 85, 373–383. 39. Crowe, S., Mills, J., McGrath, M. S. (1987) Quantitative immunocytofluorographic analysis of CD4 surface antigen expression and HIV infection of human peripheral blood monocyte/macrophages. AIDS Res. Hum. Retroviruses 3, 135–145. 40. Kedzierska, K., Maerz, A., Warby, T., Jaworowski, A., Chan, H., Mak, J., Sonza, S., Lopez, A., Crowe, S. (2000) Granulocyte-macrophage colonystimulating factor inhibits HIV-1 replication in monocyte-derived macrophages. AIDS 14, 1739 –1748. 41. Li, X., Ward, C., Thien, F., Bish, R., Bamford, T., Bao, X., Bailey, M., Wilson, J. W., Haydn Walters, E. (1999) An antiinflammatory effect of salmeterol, a long-acting ␤(2) agonist, assessed in airway biopsies and bronchoalveolar lavage in asthma. Am. J. Respir. Crit. Care Med. 160, 1493–1499. 42. Kedzierska, K., Vardaxis, N. J., Jaworowski, A., Crowe, S. M. (2001) Fc␥R-mediated phagocytosis by human macrophages involves Hck, Syk, and Pyk2 and is augmented by GM-CSF. J. Leukoc. Biol. 70, 322–328. 43. Kedzierska, K., Ellery, P., Mak, J., Lewin, S. R., Crowe, S. M., Jaworowski, A. (2002) HIV-1 down-modulates ␥ signaling chain of Fc ␥ R in human macrophages: a possible mechanism for inhibition of phagocytosis. J. Immunol. 168, 2895–2903. 44. Ziegler-Heitbrock, H. W. (1996) Heterogeneity of human blood monocytes: the CD14⫹ CD16⫹ subpopulation. Immunol. Today 17, 424 – 428. 45. Levy, P. C., Utell, M. J., Fleit, H. B., Roberts Jr., N. J., Ryan, D. H., Looney, R. J. (1991) Characterization of human alveolar macrophage Fc ␥ receptor III: a transmembrane glycoprotein that is shed under in vitro culture conditions. Am. J. Respir. Cell Mol. Biol. 5, 307–314.

302


46. Levine, S. J. (2004) Mechanisms of soluble cytokine receptor generation. J. Immunol. 173, 5343–5348. 47. Arribas, J., Merlos-Suarez, A. (2003) Shedding of plasma membrane proteins. Curr. Top. Dev. Biol. 54, 125–144. 48. Arend, W. P., Ammons, J. T., Kotzin, B. L. (1987) Lipopolysaccharide and interleukin 1 inhibit interferon-␥-induced Fc receptor expression on human monocytes. J. Immunol. 139, 1873–1879. 49. Liao, G., Simon, S. R. (1994) Temporal down-regulation of Fc ␥ RIII expression and Fc ␥ receptor-mediated phagocytosis in human monocytederived macrophages induced by TNF-␣ and IL-1 ␤. J. Leukoc. Biol. 55, 702–710. 50. Tridandapani, S., Wardrop, R., Baran, C. P., Wang, Y., Opalek, J. M., Caligiuri, M. A., Marsh, C. B. (2003) TGF-␤ 1 suppresses [correction of supresses] myeloid Fc ␥ receptor function by regulating the expression and function of the common ␥-subunit. J. Immunol. 170, 4572– 4577. 51. Wahl, S. M., Allen, J. B., Welch, G. R., Wong, H. L. (1992) Transforming growth factor-␤ in synovial fluids modulates Fc ␥ RII (CD16) expression on mononuclear phagocytes. J. Immunol. 148, 485– 490. 52. Welch, G. R., Wong, H. L., Wahl, S. M. (1990) Selective induction of Fc ␥ RIII on human monocytes by transforming growth factor-␤. J. Immunol. 144, 3444 –3448. 53. Randolph, G. J., Sanchez-Schmitz, G., Liebman, R. M., Schakel, K. (2002) The CD16(⫹) (Fc␥RIII(⫹)) subset of human monocytes preferentially becomes migratory dendritic cells in a model tissue setting. J. Exp. Med. 196, 517–527. 54. Middelhoven, P. J., Van Buul, J. D., Hordijk, P. L., Roos, D. (2001) Different proteolytic mechanisms involved in Fc ␥ RIIIb shedding from human neutrophils. Clin. Exp. Immunol. 125, 169 –175. 55. Moldovan, I., Galon, J., Maridonneau-Parini, I., Roman Roman, S., Mathiot, C., Fridman, W. H., Sautes-Fridman, C. (1999) Regulation of production of soluble Fc ␥ receptors type III in normal and pathological conditions. Immunol. Lett. 68, 125–134. 56. Ravetch, J. V., Perussia, B. (1989) Alternative membrane forms of Fc ␥ RIII(CD16) on human natural killer cells and neutrophils. Cell typespecific expression of two genes that differ in single nucleotide substitutions. J. Exp. Med. 170, 481– 497. 57. Huizinga, T. W., de Haas, M., Kleijer, M., Nuijens, J. H., Roos, D., von dem Borne, A. E. (1990) Soluble Fc ␥ receptor III in human plasma originates from release by neutrophils. J. Clin. Invest. 86, 416 – 423. 58. Fischer-Smith, T., Croul, S., Sverstiuk, A. E., Capini, C., L’Heureux, D., Regulier, E. G., Richardson, M. W., Amini, S., Morgello, S., Khalili, K., Rappaport, J. (2001) CNS invasion by CD14⫹/CD16⫹ peripheral bloodderived monocytes in HIV dementia: perivascular accumulation and reservoir of HIV infection. J. Neurovirol. 7, 528 –541. 59. Sonza, S., Mutimer, H. P., Oelrichs, R., Jardine, D., Harvey, K., Dunne, A., Purcell, D. F., Birch, C., Crowe, S. M. (2001) Monocytes harbor replication-competent, non-latent HIV-1 in patients on highly active antiretroviral therapy. AIDS 15, 17–22. 60. Crowe, S., Zhu, T., Muller, W. A. (2003) The contribution of monocyte infection and trafficking to viral persistence, and maintenance of the viral reservoir in HIV infection. J. Leukoc. Biol. 74, 635– 641. 61. Orenstein, J. M., Fox, C., Wahl, S. M. (1997) Macrophages as a source of HIV during opportunistic infections. Science 276, 1857–1861. 62. Igarashi, T., Brown, C. R., Endo, Y., Buckler-White, A., Plishka, R., Bischofberger, N., Hirsch, V., Martin, M. A. (2001) Macrophage are the principal reservoir and sustain high virus loads in rhesus macaques after the depletion of CD4⫹ T cells by a highly pathogenic simian immunodeficiency virus/HIV type 1 chimera (SHIV): implications for HIV-1 infections of humans. Proc. Natl. Acad. Sci. USA 98, 658 – 663. 63. Gavin, A. L., Wines, B. D., Powell, M. S., Hogarth, P. M. (1995) Recombinant soluble Fc ␥ RII inhibits immune complex precipitation. Clin. Exp. Immunol. 102, 620 – 625. 64. Wines, B. D., Gavin, A., Powell, M. S., Steinitz, M., Buchanan, R. R., Mark Hogarth, P. (2003) Soluble Fc␥RIIa inhibits rheumatoid factor binding to immune complexes. Immunology 109, 246 –254.

http://www.jleukbio.org