Uptake of Leishmania major by dendritic cells is mediated by Fc receptors and facilitates acquisition of protective immunity

http://www.100md.com 《实验药学杂志》

     1 Department of Dermatology and 2 Section for Pathophysiology, First Department of Internal Medicine, Johannes Gutenberg-University, Mainz 55131, Germany

    3 Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases and 4 Dermatology Branch, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD 20892

    5 Department of Dermatology, University of Münster, Münster 48129, Germany

    6 Department of Human and Clinical Genetics, Leiden University Medical Center, Leiden 2300, Netherlands

    CORRESPONDENCE Esther von Stebut: vonstebu@mail.uni-mainz.de

    Uptake of Leishmania major by dendritic cells (DCs) results in activation and interleukin (IL)-12 release. Infected DCs efficiently stimulate CD4– and CD8– T cells and vaccinate against leishmaniasis. In contrast, complement receptor 3–dependent phagocytosis of L. major by macrophages (M) leads exclusively to MHC class II–restricted antigen presentation to primed, but not naive, T cells, and no IL-12 production. Herein, we demonstrate that uptake of L. major by DCs required parasite-reactive immunoglobulin (Ig)G and involved FcRI and FcRIII. In vivo, DC infiltration of L. major–infected skin lesions coincided with the appearance of antibodies in sera. Skin of infected B cell–deficient mice and Fc–/– mice contained fewer parasite-infected DCs in vivo. Infected B cell–deficient mice as well as Fc–/– mice (all on the C57BL/6 background) showed similarly increased disease susceptibility as assessed by lesion volumes and parasite burdens. The B cell–deficient mice displayed impaired T cell priming and dramatically reduced IFN- production, and these deficits were normalized by infection with IgG-opsonized parasites. These data demonstrate that DC and M use different receptors to recognize and ingest L. major with different outcomes, and indicate that B cell–derived, parasite-reactive IgG and DC FcRI and FcRIII are essential for optimal development of protective immunity.

    Abbreviations used: BMDC, bone marrow–derived DC; CFSE, carboxyl fluorescein succinimidyl ester; CR3, complement receptor 3; IS, immune serum; M, macrophages; NMS, normal mouse serum; SLA, soluble Leishmania antigen.

    F. Woelbing and S.L. Kostka contributed equally to this paper.

    In cutaneous leishmaniasis affecting mice and man, control of infection is associated with Th1/Tc1-mediated, IFN-–dependent elimination of intracellular parasites (1, 2). After infection of mice with physiologic low dose inocula of Leishmania major parasites, the evolution of skin lesions occurs in three distinct phases (3). In the initial "silent" phase, resident macrophages (M) phagocytose L. major promastigotes primarily via complement receptor 3 (CR3) (4, 5), which inactivates the infected cells and allows parasite amplification (as intracellular amastigotes) at sites of infection. In the second phase, development of clinically evident lesions occurs coincident with the influx of inflammatory cells, including neutrophils, M, and eosinophils. Subsequently, immunity is initiated by infiltration of DCs as well as T and B cells, and lesions resolve (the third phase) (3).

    Both M and DCs, the major APCs in skin, clearly influence the development of cellular immune responses against Leishmania. Dermal M capture organisms at sites of inoculation and, after establishment of protective immunity, they ultimately kill the parasites. However, M do not actively participate in T cell priming. In all likelihood, DCs take up amastigotes of L. major present in lesional skin, become activated, and migrate to draining LN where they present Leishmania antigen to naive T cells (6, 7). There are striking differences in the ways that M and DCs interact with L. major parasites in vitro. First, skin DCs preferentially take up L. major amastigotes, the obligate intracellular life form of the parasite, rather than promastigotes (transmitted by sand flies), whereas M efficiently phagocytose both life forms (7–9). Second, the phagocytotic capacity of DCs is limited with regard to efficiency and capacity as compared with that of M (7). Third, L. major–infected DCs, unlike infected M, release IL-12 and efficiently induce Th1/Tc1 differentiation of naive cells (7, 10–12). Fourth, although both cell types present Leishmania antigen via the MHC class II pathway, only DCs prime and restimulate L. major–specific CD8+ T cells (13).

    Based on the different behaviors and functional roles of M and DCs in L. major infections, we hypothesized that DCs and M might take up L. major via different phagocytotic receptors. M ingest L. major via CR3-dependent mechanisms (4). Herein, we identified immune IgG and Fc receptors (FcRI and FcRIII) as critical mediators of L. major uptake by DCs in vitro. In vivo, mice infected with IgG-opsonized parasites showed enhanced protective immunity as well as increased numbers of L. major–infected lesional DCs. We also determined that B cell– (μMT and JHT) and Fc-deficient mice had decreased numbers of L. major–infected lesional DCs and enhanced lesion progression. In addition, we observed impaired CD4- and CD8-priming in the absence of B cells. Immune IgG production and engagement of DC FcR are required for timely development of Th1/Tc1-dependent immunity and control of experimental cutaneous leishmaniasis in mice.

    RESULTS

    CR3 does not mediate uptake of L. major by DCs

    M phagocytosis of L. major promastigotes and amastigotes is rapid and efficient (1). In contrast, skin DCs preferentially ingest amastigotes, and this occurs slowly and inefficiently (7). We generated bone marrow–derived DCs (BMDCs) using GM-CSF/IL-4 and confirmed our previous findings obtained with skin DCs. Day 6 immature DCs expressed CD11c, intermediate levels of MHC class II, and low levels of CD86 (Fig. 1 A). BMDCs, like skin DCs, internalized freshly isolated amastigotes in a time- and dose-dependent manner. Normal mouse serum (NMS)-opsonized promastigotes, in contrast, were not readily ingested (27 ± 6 vs. 8 ± 1% infected DCs with a DC/parasite ratio of 1:3 at 18 h; P 0.05, Fig. 1 B). As expected, DC infection was associated with up-regulation of MHC class I/II and costimulatory markers (reference 7 and unpublished data).

    Figure 1. L. major amastigotes, rather than promastigotes, are preferentially internalized by DCs independent from CR3/CR4. Bone marrow–derived DCs and amastigotes or promastigotes of L. major were cocultured at various ratios at 2 x 105 DCs/ml. (A) Before coculture, surface phenotypes of immature DCs were verified by FACS. (B) At the indicated time points, cells were harvested, cytospun, and the percentage of infected cells was determined (mean ± SEM, n 3, *, P 0.05, **, P 0.005, ***, P 0.002). (C) DCs from CD18–/– and wild-type 129 x C57BL/6 controls were cocultured with L. major (1:3). (D) C57BL/6 DC were preincubated with 5 mg/ml mannan, 50 μg/ml anti-CD11b, anti-DEC205, or isotype control before amastigotes of L. major were added (1:3). (C and D) After 18 h, cells were harvested and cytospins were analyzed for the percentage of infected DC (mean ± SEM, n 3, *, P 0.05).

    Phagocytosis of L. major by M is CR3 dependent (5). To investigate the role of CR3 and CR4 in L. major uptake by DCs, we used CD18–/– mice. As expected, DCs generated from CD18–/– mice did not express CD11b or CD11c (unpublished data). No differences in the percentages of infected wild type or CD18–/– DCs (Fig. 1 C) or the number of parasites/cell was observed after DCs and L. major amastigotes were cocultured for 18 h.

    We also assessed the involvement of other candidate receptors. Antibodies reactive with CD11b (clone M1/70) (9), CD205 (clone NLDC145) (14), or preincubation with mannan (5) were used at optimal concentrations. This concentration of mannan was able to completely inhibit the uptake of C. albicans by M (unpublished data) (5). None of the inhibitors tested affected the uptake of L. major by DCs (Fig. 1 D). Thus, CR3/CR4 and C-type lectins appear to be dispensable for phagocytosis of L. major by DCs.

    Immunoglobulins enhance uptake of L. major by DCs

    L. major amastigotes are isolated from infected tissues, whereas metacyclic promastigotes are enriched from stationary phase in vitro cultures. Among the most prominent differences between surface characteristics is the large amount of Ig bound to the surfaces of amastigotes, but not promastigotes. To determine if Ig was involved in parasite uptake, we quantified the ability of amastigotes isolated from B cell–replete, wild-type BALB/c mice, μMT (B cell–deficient) mice and SCID (B cell– and T cell–deficient) mice to parasitize DCs. DCs readily phagocytosed amastigotes from BALB/c mice, but not amastigotes from μMT or SCID mice (Fig. 2, A and B). Opsonization with NMS did not affect uptake. Parasites from B cell–deficient mice efficiently entered DCs only after they had been preincubated with Ig-containing immune serum (IS) from L. major–infected BALB/c mice (or C57BL/6 mice; unpublished data). Phagocytosis of amastigotes by M was not affected by the presence or absence of Ig. Opsonization of amastigotes from B cell–deficient mice with IS (Fig. 2 C) also induced enhanced release of IL-12p40 from DCs, whereas infection of M did not promote IL-12 production. Ig-mediated uptake of amastigotes did induce IL-10 release from M (15), whereas little, if any, IL-10 was produced by infected DCs.

    Figure 2. Immunoglobulins are required for internalization of L. major amastigotes by DCs. Amastigotes of L. major were prepared from BALB/c mice or mice deficient for B cells (μMT or SCID mice). As indicated, parasites were opsonized for 10 min with 5% of normal mouse serum (NMS) or serum from 6 wk L. major–infected BALB/c mice (IS) and added to DCs or skin-M from C57BL/6 mice (2 x 105 cells/ml) at a parasite/cell ratio of 3:1. (A and B) 18 h later, cells were harvested and cytospun and the percentage of infected cells was determined by light microscopy and expressed as mean ± SEM (n 4, *, P 0.05, ***, P 0.002). (C) The coculture supernatants were assayed for the presence of IL-12p40 and IL-10 by ELISA (mean ± SEM, n 4).

    To determine if Ig-coated promastigotes could be ingested by DCs, metacyclic L. major promastigotes were left untreated or were opsonized with NMS or IS for 10 min at 37°C. After washing, parasites were cocultured with DCs for 18 h. IS-treated promastigotes were efficiently taken up by DCs and induced IL-12 release, whereas untreated or NMS-treated parasites were not ingested (Fig. 3 A). Interestingly, complete transformation of promastigotes into amastigotes was not observed within all DCs, even after 18 h (Fig. 3 B), suggesting that there are differences in the phagosomal compartments of DCs and M that influence this transition (e.g., differences in pH, content of proteolytic enzymes) (16). Stimulation of antigen-specific, carboxyl fluorescein succinimidyl ester (CFSE)-labeled T cells with parasite-treated DCs revealed that DCs infected with NMS amastigotes or IS promastigotes induced similar expansion of both CD4+ and CD8+ T cells, whereas DCs treated with NMS promastigotes did not promote T cell proliferation (Fig. 3 C).

    Figure 3. Infectious stage promastigotes of L. major infect DCs in the presence of IgG. DCs were cocultured with metacyclic promastigotes of L. major that had either been left untreated or were opsonized with normal mouse serum (NMS) or serum of 6 wk–infected BALB/c mice (IS) before addition to the cultures. (A and B) Cells were harvested after 18 h and infection rates were determined on cytospins. The IL-12p40 production of DCs was analyzed by ELISA (mean ± SEM, n = 3, *, P 0.05; **, P 0.005). (C) BMDCs infected as indicated were coincubated (1:2) with CFSE-labeled LN cells (at 106 cells/200 μl) obtained from 6 wk–infected C57BL/6 mice. Antigen-specific expansion of CD4+ and CD8+ T cells was assessed and the relative number of Leishmania-reactive cells (in percents of total CD4+ or CD8+ T cells) was determined in DC-stimulated cultures compared with untreated control cultures. One representative FACS result is shown on the left, pooled data is shown on the right (mean ± SEM, n = 3, *, P 0.05). (D) Parasites were analyzed for surface binding of Ig subclasses after different opsonization procedures. Amastigotes were isolated from infected C57BL/6 tissue and opsonized. Promastigotes were enriched from stationary phase cultures and opsonized similarly. One out of three independent experiments with similar results is shown. (E) IgG was purified from IS. IgG as well as the IgGneg fraction were used for opsonization of promastigotes. Opsonized promastigote preparations were cocultured with DCs for 18 h at a ratio of 3:1 (2 x 105 cells/ml). Cells were harvested and the percentage of infected cells was determined on cytospins (mean ± SEM, n = 3, *, P 0.05).

    Phagocytosis of L. major is IgG dependent

    Amastigotes from infected tissue efficiently parasitize DCs. By flow cytometry, we detected both IgG1 and IgG2a/b on the surfaces of amastigotes when analyzed directly after isolation from C57BL/6 mice or after opsonization with NMS (Fig. 3 D). Additional opsonization with NMS or IS led to enhanced binding of IgM to amastigotes. Cell surface Ig was not detected on promastigotes. NMS-opsonized promastigotes exhibited surface-associated IgM, attributable to natural Ig found in the sera of naive mice (17). Because both NMS- and IS-opsonized amastigotes were taken up to similar extents and NMS-opsonized promastigotes were not phagocytosed by DCs (compare with Fig. 3 A), we conclude that IgM is not required for parasite uptake. Interestingly, promastigotes bound similar amounts of IgG1 and IgG2a/b when incubated with sera harvested from infected resistant C57BL/6 mice (Fig. 3 D) or susceptible BALB/c mice (unpublished data).

    To conclusively implicate immune IgG in DC-parasite uptake, we isolated total IgG from IS using protein G affinity columns and tested the capacity of IgG to trigger phagocytosis. Similar to IS, the IgG fraction mediated uptake of promastigotes, whereas parasites incubated with the IgG-depleted fraction were not phagocytosed (Fig. 3 E). In addition, parasite uptake was associated with IL-12p40 release (772 ± 324 pg/ml for DCs incubated with IgG promastigotes vs. 137 ± 27 pg/ml for DCs cocultured with promastigotes incubated with the IgGneg fraction; n 3, P = 0.03).

    Both FcRI and FcRIII mediate uptake of L. major by DCs

    IgG1-containing immune complexes bind preferentially to FcRIII (and FcRII) and IgG2a-containing complexes bind with higher affinity to FcRI than to FcRIII. FcRII typically mediates endocytosis of soluble immune complexes (18). DCs from knockout mice deficient for single FcR family members ingested L. major as efficiently as DCs from wild-type mice (Fig. 4 A). In addition, blocking antibodies directed against FcRII/III (clone 2.4G2) did not have a dramatic effect on L. major uptake by wild-type DCs (Fig. 4 B, left). However, significant inhibition of L. major phagocytosis by DCs (up to 70%) was observed if DCs from FcRI/III- or Fc-deficient mice were compared with wild-type cells (Fig. 4 B). Uptake of amastigotes and Ig-opsonized promastigotes was impaired to similar extents. Thus, FcRI and FcRIII each facilitate phagocytosis of L. major by DCs, and these receptors can compensate for one another.

    Figure 4. Fc RI/III mediate uptake of L. major parasites by DCs. Bone marrow–derived DCs from C57BL/6, FcRII–/–, FcRI–/–, FcRIII–/–, Fc–/–, and FcRI/III–/– were cocultured with L. major at a ratio of 1:3 (2 x 105 cells/ml). Parasites (Am = amastigotes, Prom = promastigotes) were opsonized with normal mouse serum (NMS) or serum of 6 wk–infected BALB/c mice (IS) for 10 min before addition to the cultures. As indicated, 50 μg/ml anti-CD16/32 (clone 2.4G2) or isotype control was added to the cells 1 h before the parasites (B). Cells were harvested after 18 h. The percentage of infected cells was determined (mean ± SEM, n 3, *, P 0.05, **, P 0.005 compared with isotype treatment or wild type control ).

    Accumulation of infected DCs in lesions coincides with the appearance of Leishmania-specific IgG in sera

    In the setting of physiologic low dose infections, we have shown that increased accumulation of both T cells and DCs at inoculation sites coincides with the onset of lesion involution (3). In addition, infiltration with DCs was delayed as compared with M recruitment and infection. DCs were identified in lesions beginning 5 wk after inoculation, and their number increased substantially during the healing phase.

    To determine if immune IgG, which dramatically enhances L. major infection of DCs in vitro, is present at the time that DCs are recruited to Leishmania lesions, we infected C57BL/6 mice with 103 promastigotes and quantified the number of inflammatory cells in lesional skin as well as the appearance of Leishmania-reactive IgG in sera at weekly intervals. Fig. 5 shows that by weeks 5–6 after infection, the numbers of DCs as well as serum parasite-specific IgG levels were increased. This indicates that Leishmania-specific IgG is available to opsonize parasites and enhance phagocytosis by DCs at the time that DCs are infected in vivo. Significant accumulation of CD19+ B cells in lesional skin (>103 cells) was not detected within 8 wk after infection.

    Figure 5. The appearance of parasite-specific IgG is coincident with DCs recruitment into Leishmania lesions. Accumulation of DCs into skin lesions of L. major–infected C57BL/6 mice (103 metacyclic promastigotes intradermally, day 0) was determined as described previously. In parallel, the anti-Leishmania antibody response was monitored in the serum by ELISA (mean ± SEM, n = 5 mice). One out of two independent experiments with similar results is shown.

    In vivo targeting of DCs with IgG-opsonized promastigotes speeds disease resolution

    Previously, we and others have demonstrated that L. major–infected DCs release IL-12 and effectively vaccinate against progressive disease (10–12). Therefore, infection of DCs in vivo earlier in the course of infection should accelerate development of Th1 immunity. To test this hypothesis, promastigotes were opsonized either with NMS or IS. After washing, low dose infections using 103 opsonized parasites were initiated in the ear skin of C57BL/6 mice (Fig. 6). Inflammatory dermal cells from lesional ear skin were studied weekly (Fig. 6, A and B). Interestingly, the numbers of CD11c+ DCs in IS promastigote–infected skin were significantly higher (7.9 ± 0.9 x 104/lesion) than those in NMS promastigote–treated ears (2.5 ± 0.5 x 104/lesion in week 1, n = 3, P 0.005), especially at early time points. At later time points (week 3 and after), this difference was not evident. DCs were enriched by preparative flow sorting and the number of infected DCs was determined (Fig. 6, B and C). At early time points, the percentage of DCs containing intracellular amastigotes was low. However, by week 2, significantly more infected cells were found in ears of mice infected with IS- versus NMS-opsonized parasites (9.5 ± 1.3 vs. 3.4 ± 0.7%, n = 3, P 0.002). By week 3, this difference also disappeared.

    Figure 6. Increased recruitment and infection of DCs after injection of IgG-opsonized promastigotes of L. major leads to enhanced protection in vivo. C57BL/6 mice were infected intradermally into ear skin with 103 metacyclic promastigotes that have been opsonized with normal mouse serum (NMS) or serum from 6 wk–infected BALB/c mice (IS) and washed. Inflammatory ear cells were isolated and analyzed for the presence of CD11c+ DCs at different time points by flow cytometry (A). The number of DCs per ear was calculated (mean ± SEM, n = 3, **, P 0.005; B). CD11c+ DCs were purified using flow sorting and cytospun. The percentage of infected DCs was determined by light microscopy (mean ± SEM, n = 3, ***, P 0.002; B and C). (D) Lesion development was monitored over the course of >3 mo. Lesion volumes are shown as mean ± SEM (**, P 0.005 and ***, P 0.002, n 10 mice). (E) Parasite loads of infected mice were determined at the indicated time points using limiting dilution assays. Each data point represents the number of organisms from one ear and the bars indicate arithmetic means. One representative out of two independent experiments is shown. (F) Cytokine levels were determined in LN cultures stimulated in the presence of soluble Leishmania antigen by ELISA (mean ± SEM, *, P 0.05, n 10).

    Lesion development in infected mice was monitored for >3 mo. Interestingly, cutaneous lesions of mice infected with IS-opsonized parasites were significantly smaller and resolved more quickly than those in mice that were infected with parasites opsonized with NMS (Fig. 6 D). In addition, lesional parasite loads were decreased in weeks 4 and 6 after infection in mice inoculated with IS-opsonized parasites compared with NMS-treated parasites (Fig. 6 E). Smaller lesion volumes were associated with increased Th1 immunity as measured by antigen-specific restimulation of LN cells at weeks 4 and 6 (Fig. 6 F). The IFN-/IL-4 ratio in IgG-parasite infected mice was Th1-predominant (week 6: 1,068 ± 250) as compared with mice infected with NMS-opsonized parasites (week 6: 382 ± 86, n = 6, P 0.05). Collectively, these data suggest that enhanced IgG-mediated recruitment and L. major infection of DCs in vivo leads to enhanced Th1 immunity and more rapid resolution of cutaneous lesions.

    B cell–deficient mice show enhanced lesion progression associated with decreased numbers of infected DCs and impaired CD4- and CD8-priming

    Because our data suggested that IgG mediates parasite uptake by DCs, we characterized L. major infections in B cell–deficient μMT mice (19). Herein, wild-type C57BL/6 or μMT mice were infected with physiologically relevant doses of L. major (103 promastigotes). Compared with wild-type mice, μMT mice showed significantly enhanced lesion progression from week 6 after infection (Fig. 7 A). Lesion involution was delayed by 4 wk in μMT compared with control mice. Furthermore, the skin of μMT mice contained greater numbers of parasites reaching a peak load of 4 ± 2 x 105 parasites/ear at week 6 as compared with 3 ± 2 x 104 parasites/ear in wild types (P 0.05) (Fig. 7 B). The IFN-/IL-4 ratios of μMT LN cell cultures stimulated with soluble Leishmania antigen (SLA) were also skewed toward a Th2 profile as compared with C57BL/6 cells. In weeks 6 and 8 after infection, μMT LN cells released significantly less IFN- and more IL-4 compared with C57BL/6 mice (e.g., 40.1 ± 12.6 in μMT compared with 100.7 ± 19.2 ng IFN-/ml in C57BL/6 mice in week 6, n 9, P 0.05; Fig. 7 C).

    Figure 7. Increased lesion progression in μMT mice due to decreased numbers of L. major–infected lesional DC and impaired T cell priming. Groups of 5 C57BL/6 or B cell–deficient μMT mice were infected into ear skin with 103 metacyclic promastigotes. (A) Lesion development was monitored over the course of >3 mo (mean ± SEM, *, P 0.05, **, P 0.005, and ***, P 0.002, n 3). (B) Lesional parasite loads were determined; bars indicate arithmetic means. Pooled data of two to three experiments are shown. (C) LN cells were harvested and antigen-specific cytokine release was determined after 48 h using ELISA specific for murine IFN- and IL-4 (mean ± SEM, n 5, *, P 0.05). (D) Inflammatory ear cells were isolated and CD11c+ DCs were purified using flow sorting. The percentage of infected DCs was determined on cytospins (mean ± SEM, n = 3, *, P 0.05). (E and F) Antigen-specific proliferation of CD4+ and CD8+ T cells was determined in week 6 after infections. LN cells were labeled with CFSE and subcultured in the presence of soluble Leishmania antigen (SLA). T cells were pregated using staining for CD4 or CD8. For each mouse, the relative number of Leishmania-reactive cells (in percentage of total CD4+ or CD8+ T cells) was calculated in antigen-stimulated compared with untreated control cultures (mean ± SEM, n = 2, *, P 0.05).

    We also isolated inflammatory cells from infected ears of μMT and wild-type mice. No significant difference in the number of CD11c+ DCs that accumulated in the lesions of μMT mice as compared with C57BL/6 ears was found (unpublished data). However, lesions of μMT mice contained significantly fewer L. major–infected DCs at several time points (Fig. 7 D). Finally, we sought to determine if there was a correlation between numbers of infected DCs and the ability to prime CD4 and CD8 T cells in situ. LN cells of infected C57BL/6 or μMT mice were isolated 6 wk after infection and labeled with CFSE. Antigen-specific expansion of CD4+ and CD8+ T cells was assessed 5 d after restimulation of LN cells with SLA. μMT LN cells exhibited decreased SLA-specific CD4 expansion as compared with C57BL/6 cells (3.7 ± 1% vs. 15.7 ± 3%; Fig. 7, E and F). Interestingly, the number of Leishmania-reactive CD8+ T cells was also greatly reduced in the absence of B cells (SLA: 2.9 ± 0.5% compared with 14.1 ± 3.9% in C57BL/6 mice, n = 5, P 0.05). In summary, enhanced lesion progression in the μMT mice was associated with decreased numbers of infected DCs and defective T cell priming.

    Infection of μMT mice with IgG-opsonized parasites normalizes lesion development

    To investigate whether the deficiency in B cells or the lack of antibody contributed to the phenotype of μMT mice, we infected μMT mice with 103 NMS- or IS-opsonized promastigotes. In this setting, μMT mice infected with L. major developed lesions in the presence of immune IgG that were significantly smaller than those caused by NMS-opsonized parasites (Fig. 8 A). In parallel, decreased lesion volumes in IgG-opsonized parasite-infected μMT mice correlated with significantly smaller parasite burdens in week 6 (Fig. 8 B). In IS parasite–infected μMT mice, the IFN/IL-4 ratio was shifted from a Th2-predominant (828 ± 94) to a Th1 immune response (3,680 ± 1,515, n = 4, week 6). Thus, the lack of host IgG is responsible for disease outcome in μMT mice. The skin of μMT mice infected with NMS-opsonized or IS-opsonized promastigotes was analyzed for the presence of infected CD11c+ DC (Fig. 8 C). As shown before, infection of maximally 5% of DCs was found in μMT mice infected with NMS-treated parasites. Interestingly, inoculation of IgG-containing parasites led to dramatically increased numbers of infected DCs in the early course of infection (Fig. 8 C), even higher than those found in wild types (compare with Fig. 7 D).

    Figure 8. Normalization of cutaneous leishmaniasis in B cell–deficient μMT infected with IgG-opsonized promastigotes of L. major. Groups of 5 μMT mice were infected into ear skin with 103 NMS- or IS-opsonized metacyclic promastigotes. (A) Lesion volumes are presented as mean ± SEM (n = 3, *, P 0.05, and ***, P 0.002). (B) Parasite loads of infected ears were determined in week 6 after infection using limiting dilution assays. Each data point represents the number of organisms from one ear and bars indicate arithmetic means. One representative out of two independent experiments is shown. (C) Lesional CD11c+ DCs were purified using flow sorting at the indicated time points and the percentage of infected DCs was determined on cytospins (mean ± SEM, n = 2, *, P 0.05). (D) B cell–deficient JHT mice were infected with 103 metacyclic promastigotes and developing lesions monitored every week (mean ± SEM, *, P 0.05, and ***, P 0.002, n 10 mice/group). (E) Groups of 5 C57BL/6 Fc–/– mice were infected with 103 metacyclic promastigotes. Lesion volumes were assessed for >3 mo and are presented as mean ± SEM (*, P 0.05 and ***, P 0.002, n = 2). Parasite loads were determined in week 4 by limiting dilution assay. One representative out of two independent experiments is shown (*, P 0.05). The number of infected lesional DCs was determined on cytospins in week 4 (n = 2).

    The μMT mice were previously shown to contain Ig in the sera, at least when mice were of BALB/c genetic background (20, 21). The presence of soluble Ig is due to low-level leakiness of the locus (21). To confirm critical experiments in a truly B cell–deficient mouse strain, we infected C57BL/6 JHT mice characterized by deletion of the Ig heavy chain (22). As shown in Fig. 8 D, increased lesion development was observed in JHT mice over the course of 4 wk identical to the course of infections in μMT mice. This is in contrast with the findings of Miles et al., who reported that JH mice on a BALB/c background were less susceptible to infection than their controls (23).

    Enhanced lesion progression and decreased numbers of infected DCs in vivo in mice lacking Fc receptors

    Our data suggested that FcR-mediated uptake of L. major parasites by DCs mediates protection. Thus, we infected Fc chain–deficient mice lacking all three known activating FcR with physiologically low dose inocula of L. major (Fig. 8 E). Lesions were monitored for >3 mo. Fc–/– C57BL/6 mice developed more progressive lesions between weeks 4 and 9 as compared with wild-type controls. Maximum lesion sizes in Fc–/– mice were detected in week 9, reaching 21 ± 2 mm3 (C57BL/6: 13 ± 1 mm3, n = 14, P 0.008). Increased lesion volumes were paralleled by significantly higher parasite burdens as determined in week 4 after infection (Fig. 8 E). Similar to the course of disease in B cell–deficient mice, lesion involution in Fc–/– mice was normal and all mice ultimately healed their infection. This data suggests that FcR-mediated antibody effects are not an absolute requirement for healing.

    Finally, we assessed the number of parasite-containing CD11c+ DC in lesions of Fc–/– mice infected for 4 wk with low doses of L. major (Fig. 8 E). Ear skin of FcR-deficient mice harbored fewer parasite-infected DC (10.5 ± 2.3%) as compared with wild-type DCs (20.2 ± 3.8%, n = 4, P = 0.09). This finding confirmed our in vitro data obtained with BMDCs generated from Fc–/– mice that demonstrated inhibited parasite uptake in cocultures with L. major (Fig. 4 B).

    DISCUSSION

    Microbe-binding receptors orchestrate events that occur subsequent to phagocytosis by transducing specific cellular signals (24). The main receptor for uptake of Leishmania promastigotes by M is CR3 (4, 5). In the initial stages of cutaneous leishmaniasis, most parasites are taken up by M. CR3-mediated phagocytosis of Leishmania by M leads to selective inhibition of IL-12 release (5, 25–27). Production of IL-12 in leishmaniasis is delayed (3), and we and others have suggested that DCs, rather than M, are the primary source of this Th1-promoting cytokine. It has also been demonstrated that infected DCs are activated and effectively present L. major antigen to both naive CD4+ and CD8+ T cells in vitro and vaccinate against leishmaniasis in vivo (7, 10, 12, 13).

    Although M and DCs are ontogenically related, their roles in initiation and propagation of immune responses against L. major are distinctly different. Uptake of L. major by DCs differed significantly from that by M with regard to kinetics as well as efficiency. Therefore, we speculated that phagocytosis of parasites by DCs might be promoted by receptors other than CR3. However, a previous report suggested that uptake of L. major amastigotes by Langerhans cells/DCs was mediated via CR3 (9). In the present study, using both blocking antibodies as well as cells deficient for CR3 and CR4 (from CD18–/– mice), we were not able to detect CR3-mediated uptake of L. major by DCs. Recently, C-type lectins (DC-SIGN, DEC-205, and Dectin-1) have also been implicated in the uptake of various pathogens by DCs (14, 28–31). We were unable to implicate mannan-binding C-type lectins in phagocytosis of L. major by murine DCs.

    In this study, we demonstrate that L. major parasites are predominantly phagocytosed by DCs via FcRI and FcRIII. In line with several studies, FcR ligation was associated with DC activation and IL-12 release (32–34). We have previously shown that DCs can cross-present Leishmania antigen to CD8+ T cells (13), whereas CR3-mediated phagocytosis by M leads exclusively to MHC class II–restricted antigen presentation. These results bear some similarity to experiments evaluating the role of FcR in antitumor immunity. In Fc–/– mice, effective cross-presentation of tumor antigens by DCs was also dependent on FcR-dependent activation (35). In addition, signaling through FcRI/III facilitated efficient restimulation of tumor-reactive T cells (36). Thus, cross-presentation of both tumor-derived and L. major–associated antigens by DCs requires FcR, and is presumably dependent on production of specific antibody as well.

    In M, ingestion of amastigotes, in contrast with CR3-phagocytosed promastigotes, appears to occur through both the FcR and CR3 (15, 37). In our work and consistent with prior findings, IgG did not play an important role in the uptake of amastigotes from SCID versus BALB/c mice by inflammatory skin M (38). Our results also confirm the finding that IgG-mediated phagocytosis of L. major by M leads to strong release of IL-10, and no IL-12 synthesis (15), which might promote parasite survival (39). Thus, FcR-mediated uptake by M and DCs has opposing roles in initiating immune responses in cutaneous leishmaniasis.

    The role of B cell–derived IgG in cutaneous leishmaniasis in vivo is not fully understood yet. Polyclonal activation of human B cells leads to the production of large amounts of parasite-specific and nonspecific Ab, particularly IgM and IgG (40). Also, amastigotes released into lesional tissue from infected and lysed M appear to be coated with antiparasite antibodies (41). In this study, we show that Leishmania-specific IgG was present in sera at the time of DC accumulation in lesions. Consistent with prior findings, intradermal infection with IgG-opsonized parasites led to enhanced early recruitment of CD11c+ DCs into the lesions (38), most likely by IgG-triggered chemokine release from M (42, 43). Administration of IgG-opsonized parasites also led to enhanced infection of DC, augmented T cell priming, and limited disease as compared with inoculation of IgG-free parasites.

    Prior data and our experiments suggest that IgG-mediated effects differ significantly, dependent on the genetic background of the mice. B cell–deficient JH BALB/c mice showed improved disease outcome after infection with supraphysiologic doses of L. pifanoi and coinjection of anti-Leishmania IgG reversed their phenotype (44). Administration of IgG at or near the time of parasite inoculation worsened disease outcome in BALB/c mice (40, 45, 46). This is consistent with studies demonstrating that FcR ligation on infected M induced IL-10 release, which in turn prevented parasite elimination and promoted disease progression (15, 23). Final proof was provided by the demonstration that anti-Leishmania IgG reconstitution of JH BALB/c mice correlated with increased IL-10 production and blocking of IL-10R prevented antibody-mediated disease exacerbation (23).

    Mice on a Leishmania-resistant background lacking functional B cells (e.g., μMT C57BL/6 mice) did not exhibit a phenotype with regard to lesion development after high dose infection with L. major (19, 37, 47, 48). However, DeKrey et al. reported that C57BL/6 μMT mice infected with high-dose inocula of L. major showed reduced IFN- production after pathogen challenge (48). In our experiment, using physiologically relevant low dose inocula, μMT as well as JHT C57BL/6 mice consistently exhibited enhanced lesion progression and delayed lesion involution, higher parasite loads, and cytokine profiles consistent with a Th2-predominant immune response as compared with C57BL/6 mice. In accordance with our in vitro data, significantly fewer infected DCs were found in lesions of μMT mice. In addition, we determined that in the absence of Leishmania IgG-mediated infection of DCs, decreased numbers of Leishmania-reactive CD4+ and CD8+ T cells developed. The defects observed in μMT mice were reversed by using IgG-opsonized parasites for infection indicating that the deficiency in Ig is responsible for worsened disease outcome in B cell–deficient mice.

    As expected from the in vitro results obtained with BMDCs generated from Fc-deficient mice, we also found decreased numbers of infected DC in Fc–/– mice paralleled by increased lesion volumes over the course of several weeks and higher parasite burdens. In contrast, in prior studies, improved disease outcome of Fc–/– mice was observed using infections with L. pifanoi or L. major (44, 49). However, the mice used were on a BALB/c background and, thus, are not comparable to those used for this study. Data generated with Leishmania-resistant mice might be more physiologically relevant in a clinical setting because the course of disease in, for example, C57BL/6 mice more closely mimics L. major infections of humans.

    In summary, we propose that the two predominant APCs in skin, M and DCs, are sequentially engaged via different pathogen recognition receptors as cutaneous leishmaniasis evolves. Although in the initial "silent" phase, L. major promastigotes are primarily phagocytosed by resident M via CR3, FcR and DCs become critically important in established infections. IgG-mediated uptake of L. major by DCs leads to IL-12 production and priming of Th1/Tc1 cells, both of which are required for efficient parasite killing by lesional M. In contrast, FcR-mediated uptake of amastigotes by M induces counterregulatory IL-10 production. This may facilitate activation of regulatory T cells, which, in turn, promotes parasite persistence and maintenance of T cell memory (39, 50). The balance between CR3 and FcR-triggered anti- and proinflammatory mechanisms involving M and DCs is critical for disease outcome. The unexpected identification of immune IgG production as a prerequisite for efficient cross-priming of Leishmania-specific Th1/Tc1 cells is intriguing. In future experiments it will be important to assess the T cell dependence of Leishmania-reactive antibody production, and to identify the APCs that are involved in B cell and, if relevant, Th priming.

    MATERIALS AND METHODS

    Animals.

    6–8-wk-old BALB/c and C57BL/6 mice were purchased from the Central Animal Facility of the University of Mainz. CD18–/– mice (51) on a mixed C57BL/6J and 129/SV background were provided by K. Scharffetter-Kochaneck (Department of Dermatology, University of Ulm, Ulm, Germany). FcRII–/– mice (52) were obtained from H. Mossmann (Max Planck-Institut für Immunbiologie, Freiburg, Germany). Mice deficient for FcRIII (53) and FcRI (54) as well as FcRI/III double deficient mice (all C57BL/6 background) were provided by S. Verbeek. C57BL/6 Fc–/– were obtained from T. Saito (RIKEN Research Center for Allergy and Immunology, Yokohama, Japan) (55) or from Taconic. B cell–deficient mice (C57BL/6 SCID, μMT, JHT) were gifts from M. Neurath, K. Steinbrink, and A. Waisman (all from University of Mainz, Mainz, Germany). All animals were housed in accordance with institutional and federal guidelines. All experiments were undertaken with approved license from the Animal Care and Use Committee of the Region Rheinland-Pfalz.

    Cells.

    Inflammatory skin-derived M (M) were elicited by subcutaneous injection of polyacrylamide beads and enriched to homogeneity (7). BMDCs were generated in GM-CSF– and IL-4–containing media (56) and harvested on day 6 of cell culture. The characteristics of the cell populations were assessed by flow cytometry using relevant surface markers. The following antibodies were used: anti–I-Ab,d/I-Ed (2G9), anti-CD11b (M1/70), anti-CD11c (HL3), anti-CD40 (3/23), anti-CD54 (3E2), anti-CD80 (1G10), anti-CD86 (GL1) (all from BD Biosciences/Becton Dickinson), anti-F4/80 (Serotec), and respective isotype control mAb.

    Parasites.

    Metacyclic promastigotes or amastigotes of L. major clone VI (MHOM/IL/80/Friedlin) were prepared as described previously (25, 57). Amastigotes were prepared from infected footpads of BALB/c or C57BL/6 mice, or mice genetically deficient in B cells (μMT, SCID) to obtain parasites devoid of Ig. Isolated parasites were opsonized with 5% NMS or serum from 6 wk–infected BALB/c or C57BL/6 mice (immune serum, IS) for 10 min (37°C) and washed before in vitro or in vivo infections. Parasites were stained for surface-associated Ig using isotype-specific secondary antibodies reactive with mouse Ig: anti-IgM (Serotec), anti-IgG1 (A85-1), and anti-IgG2a/b (R2-40, all from BD Biosciences). After staining, parasites were washed with PBS/2% BSA, fixed, and analyzed by flow cytometry. Anti-Leishmania IgG was prepared from pooled sera of 5–6-wk L. major–infected BALB/c mice using protein G columns (Pierce Chemical Co.) following the manufacturer's protocol. Sera were stored at –20°C before IgG purification. Purified IgG was stored at 4°C (0.8 mg/ml) in PBS before use.

    Phagocytosis and inhibition studies.

    Isolated cells were subcultured in medium (RPMI 1640/5% FCS) at 2 x 105/ml and parasites were added at the parasite/cell ratios indicated. In some experiments, cells were preincubated for 60 min with mannan (Sigma-Aldrich, 1 and 5 mg/ml), anti-CD11b, anti-CD16/32, anti-CD205, or control rat IgG (all at 50 μg/ml, all from BD Biosciences). Cells were harvested after several hours and cytospins were prepared. DiffQuick-stained cells were analyzed for the presence of intra- and extracellular parasites. At least 200 cells were counted per sample. Supernatants from parasite/cell cocultures were collected and assayed for the presence of IL-12p40 or IL-10 by ELISA (BD Biosciences).

    Assessment of B cell and DC infiltration and function in vivo.

    Groups of 5 C57BL/6 mice were infected intradermally in ear skin with 1,000 L. major promastigotes. At several time points, ears were harvested and the number of B cells and DCs that had accumulated at the site of infection was determined (3). In brief, ears were incubated with 2 mg/ml Liberase (Boehringer Ingelheim). After 2 h, cells were dissociated mechanically and counted and the frequency of CD19+ and CD11c+ cells was assessed using flow cytometry. In addition, serum from infected mice was obtained at several time points and stored at –20°C.

    Leishmania-specific IgG in serum was quantified by ELISA. Flat-bottom 96-well plates (Nunc) were coated overnight with 0.5 mg/100 μl of soluble freeze-thaw Leishmania lysate (SLA), blocked for 1 h with PBS/2% BSA/0.05% Tween 20, and incubated for 2 h with dilutions of sera or reference standard anti-Leishmania IgG (prepared from pooled sera of immune infected mice). Subsequently, biotinylated goat anti–mouse IgG (Caltag) was added (125 ng/ml) for 2 h at 20°C. ELISA plates were developed using commercially available ELISA kit components (BD Biosciences) and reaction products were quantified spectrophotometrically.

    In vivo infections using IgG-opsonized parasites and B cell– or Fc-deficient mice.

    C57BL/6, μMT, JHT, or Fc–/– mice were infected intradermally with 103 metacyclic L. major promastigotes. In some experiments, parasites were opsonized for 10 min with either NMS or IS and washed. Lesion development was assessed weekly in three dimensions using a caliper, and lesional volumes are reported (in mm3) as ellipsoids . Organisms present in lesional tissue were enumerated using limiting dilution assays (57). For measurement of cytokine production, 106 retroauricular LN cells/200 μl were added to 96-well plates in the presence of SLA (25 μg/ml). Antigen-specific IFN- and IL-4 production was determined after 48 h using ELISA (R&D Systems).

    At several time points, ears were harvested and inflammatory cells isolated using Liberase and mechanical disruption (3). The cells were counted and the frequencies of CD11c+ DC were determined using flow cytometry. CD11c+ cells were enriched to >98% purity using a high speed cell sorter (FACS Vantage SE System, Becton Dickinson) and cytospins were analyzed by light microscopy to estimate the number of infected DCs/ear.

    The frequency of daughter cells of proliferating antigen-reactive compared with nonproliferating LN T cells was estimated using flow cytometry (58–60). 6 wk after infection, LN cells were harvested and 5 x 106 cells/ml were labeled with 1 μM CFSE (Invitrogen). LN cells were subsequently plated at 106/200 μl media in a 96-well U-bottom plate and left untreated or stimulated with SEB (10 μg/ml; Sigma-Aldrich), or SLA (61). After 5 d, proliferation was determined using flow cytometry. T cells were selected for analysis using mAbs against CD4 (L3T4, RM4-5), CD8 (Ly2, 53–6.7), or isotype control mAb (all from BD Biosciences). For each mouse, the percentage of Leishmania-reactive cells compared with nonproliferating cells was calculated.

    Statistics.

    Statistical analysis was performed using the unpaired Student's t test.

    Acknowledgments

    The authors wish to thank Dr. K. Reifenberg and staff for excellent help in the animal care facility, Dr. H.-J. Peter (Department of Dermatology, Charité Berlin) for providing technical assistance with IgG purification, Dr. D. Sacks for helpful discussions, and Drs. K. Steinbrink and H. Jonuleit for critically reading the manuscript.

    This work was supported in part by grants from the Deutsche Forschungsgemeinschaft (DFG, SFB 490, and SFB548) to E. von Stebut and by the Intramural Program of the National Institutes of Health, National Cancer Institute, Center for Cancer Research to M.C. Udey.

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    Ronit Vogt Sionov1, Orly Cohen1, Shlomit Kfir1, Yael Zilberman2, and Eitan Yefenof1

    1 The Lautenberg Center for General and Tumor Immunology, The Hebrew University-Hadassah Medical School and 2 Department of Pharmacology, Faculty of Dental Medicine Founded by the Alpha-Omega Fraternity, 91120 Jerusalem, Israel

    CORRESPONDENCE Eitan Yefenof: yefenof@cc.huji.ac.il

    The mechanisms by which glucocorticoid receptor (GR) mediates glucocorticoid (GC)-induced apoptosis are unknown. We studied the role of mitochondrial GR in this process. Dexamethasone induces GR translocation to the mitochondria in GC-sensitive, but not in GC-resistant, T cell lines. In contrast, nuclear GR translocation occurs in all cell types. Thymic epithelial cells, which cause apoptosis of the PD1.6 T cell line in a GR-dependent manner, induce GR translocation to the mitochondria, but not to the nucleus, suggesting a role for mitochondrial GR in eliciting apoptosis. This hypothesis is corroborated by the finding that a GR variant exclusively expressed in the mitochondria elicits apoptosis of several cancer cell lines. A putative mitochondrial localization signal was defined to amino acids 558–580 of human GR, which lies within the NH2-terminal part of the ligand-binding domain. Altogether, our data show that mitochondrial and nuclear translocations of GR are differentially regulated, and that mitochondrial GR translocation correlates with susceptibility to GC-induced apoptosis.

    Abbreviations used: ALL, acute lymphoblastic leukemia; COX, cytochrome C oxidase; DBD, DNA-binding domain; Dex, dexamethasone; GC, glucocorticoid; GR, glucocorticoid receptor; GRE, glucocorticoid response element; hGR, human GR; LBD, ligand binding domain; mGR, membrane glucocorticoid receptor; MLS, mitochondrial localization signal; NLS, nuclear localization signal; PML, promyelocytic leukemia; TEC, thymic epithelial cell.

    Glucocorticoids (GCs) are commonly used for therapy of various hematologic malignancies, most notably acute lymphoblastic leukemia, multiple myeloma, and chronic lymphocytic leukemia (1, 2). The effectiveness of this approach is based on the ability of GCs to induce apoptosis of leukemic cells. Yet, the mechanisms by which GC causes apoptosis remain obscure (2, 3).

    Apoptosis by GC requires its binding to the glucocorticoid receptor (GR). Several GR isoforms have been identified (4), the GR isoform being the predominant active form. The GR protein contains two transactivation domains, a zinc-finger DNA-binding domain (DBD) and a ligand-binding domain (LBD) (5, 6). The DBD consists of two highly conserved zinc fingers, which are crucial for the binding to glucocorticoid response element (GRE) sequences. In addition, the first zinc-finger of DBD binds NFB and AP-1 (7, 8), and the second DBD zinc finger mediates receptor dimerization (9). The LBD binds GC as well as heat-shock proteins (10) and is also involved in receptor dimerization (6).

    Numerous studies indicate that in the absence of a ligand, GR is sequestered in the cytosol bound to a large heteromeric complex of heat-shock proteins and immunophilins (10) or to 14–3-3 (11). Upon ligand-binding, GR translocates to the nucleus where it transactivates and transrepresses multiple genes (3, 5). Transactivation occurs through interaction with GREs, whereas transrepression follows binding and inactivation of AP-1 and NFB. It is unknown which of these alterations in gene expression are essential for the apoptotic effect of GC.

    Some differences have been observed in the spectrum of genes affected by GC in GC-sensitive versus GC-resistant cells (12). Of note are the up-regulation of the pro-apoptotic gene Bim in GC-sensitive cells (13) and the down-regulation of the survival gene c-Myc (3). It has been suggested that the many effects of GC may shift the balance between prosurvival and proapoptotic factors, ultimately leading to cell death (2, 3). As AP-1 and NFB control several survival pathways, the interaction of GR with these factors is believed to play a major role in GC-mediated apoptosis (3, 5). This notion is supported by the observation that a GR mutant (GR-LS7) compromised in transactivation, but normal in transrepression, is as effective as the WT receptor in inducing apoptosis (14). However, a dimerization-defective GR is unable to mediate GC-induced apoptosis, although it causes transrepression through interaction with AP-1 and NFB (9). Thus, interference with AP-1 and NFB functions per se is insufficient for inducing apoptosis. The role of transrepression is further questioned by the study of Tao et al. (7), who showed that a GR mutant deficient in transrepression is still able to induce apoptosis. Because dimerization is required for DNA binding (9), it was suggested that DNA binding is a prerequisite for eliciting the apoptotic response. However, a DNA-binding defective variant of GR, which is mainly localized in the cytosol, still induces apoptosis (15). These data suggest that GC may induce apoptosis by a yet unknown pathway, which is independent of the nuclear effects of GR.

    Although GR is predominantly cytosolic, a plasma membrane form of GR (mGR) has been detected in some lymphoid cells (16, 17). Expression of mGR was found to be associated with GR mRNA transcript 1A (18), which is one of several GR transcripts expressed most abundantly in hematopoietic cell lines sensitive to GC-induced apoptosis (19, 20). It was thus suggested that mGR may be responsible for GC-induced apoptosis. Other studies, however, did not show correlation between expression of mGR and sensitivity to GC (2).

    Several reports have shown that GR translocates to the mitochondria in addition to its well-characterized nuclear translocation (21–24). As GC-induced apoptosis is mediated via the mitochondrial pathway (2), we addressed the question whether GR translocation to the mitochondria is essential for GC-induced apoptosis. We found correlation between mitochondrial GR translocation and sensitivity of cells to GC-induced apoptosis. Moreover, a GR variant that is exclusively expressed in the mitochondria enabled apoptosis of various cancer cell lines. Altogether, our data indicate a role for mitochondrial GR in eliciting apoptosis.

    RESULTS

    GC-sensitive PD1.6 and 2B4 cells do not express plasma membrane GR

    We used a series of lymphoma and leukemia cells to study the mechanisms of GC-induced apoptosis. Susceptibility to GC-induced apoptosis was determined by appearance of subdiploid cells and caspase 3 activation. PD1.6 and 2B4 cells readily apoptose in response to GC, whereas B10, S49, NB4, and Jurkat cells are resistant to GC-induced apoptosis (Fig. 1, A–C). It should be noted that the S49 variant used in this study (25) differs from the S49 variant described by Gametchu et al. (16) in being resistant to GC-induced apoptosis. Because it has been proposed that mGR is a mediator of GC-induced apoptosis (18), we compared mGR expression on GC-sensitive and resistant cells. It appears that S49, but not PD1.6, B10, 2B4, or Jurkat cells, express mGR (Fig. 1 D, subpanels A vs. C, E, G, and I, respectively). Also, the macrophage cell line RAW264.7 expresses a high level of mGR and is resistant to GC-induced apoptosis (unpublished data). Treatment with 100 nM dexamethasone (Dex) for 2 h did not induce expression of mGR on otherwise mGR-negative cells (Fig. 1 D, subpanels D, F, H, and J). The fact that GC-resistant S49 and RAW264.7 cells express mGR indicates that expression of mGR per se does not impose susceptibility to GC. The observation that GC-sensitive PD1.6 and 2B4 cells do not express mGR suggests that another mechanism is involved in this apoptotic process.

    Figure 1. (A) Sensitivity of various lymphoma and leukemia cell lines to Dex-induced apoptosis. Cells were incubated with 100 nM Dex for 20 h, and the DNA content measured by flow cytometry using propidium iodide. Percentage of subdiploid cells is given. (B) Dose–response to Dex. Cells were incubated with various concentrations of Dex and processed as in A. (C) Caspase 3 activation. Untreated or Dex-treated cells were stained for activated caspase 3 as described in Materials and methods. Percentage of positive cells is given. (D) Expression of mGR on lymphoma and leukemia cells. Untreated cells or cells treated with 100 nM Dex for 2 h were incubated with M20 antibodies to GR and FITC-conjugated goat anti–rabbit IgG (dashed line) or FITC-conjugated goat anti–rabbit IgG only (solid line). The fluorescence intensity was measured by flow cytometry.

    Translocation of GR to the mitochondria and sensitivity to GC-induced apoptosis

    It has recently been shown that GR may be localized to the mitochondria (24). As GC-evoked apoptosis is mediated via the mitochondrial pathway (2), we asked whether GR translocation to the mitochondria may be a trigger of apoptosis. To answer this question, we analyzed the intracellular trafficking of GR in GC-sensitive and resistant cells after treatment with Dex. Dex-induced GR translocation to the mitochondria was demonstrated in GC-sensitive PD1.6 cells using two different methods (Fig. 2, A, B, and E). First, immunofluorescence studies using M20 antibodies to GR and red mitotracker to visualize the mitochondria show that a certain fraction of GR localizes to the mitochondria in Dex-treated PD1.6 cells (Fig. 2 A). Second, PD1.6 cells treated with 100 nM Dex for 2 h were fractionated using the Oncogene cytosol/mitochondria fractionation kit. The quantities of GR in the cytosolic and mitochondrial fractions were analyzed by Western blotting (Fig. 2 B). The PA1-511A antibody to GR (epitope aa 346–367) resolves two major bands on Western blots. The top one is GR, whereas the bottom band is of unknown character and considered in Fig. 2 as a "background band." GR expression in the mitochondrial fraction of PD1.6 cells is increased after Dex treatment (Fig. 2 B, subpanels C and D, lane 2 vs. lane 1), indicating mitochondrial translocation of GR. Contamination of the mitochondrial fraction with mGR is excluded, because PD1.6 cells do not express mGR even after Dex treatment (Fig. 1 D, subpanels C and D). As expected, the amount of GR in the cytosolic fraction is reduced by Dex (Fig. 2 B, subpanel A, lane 2 vs. lane 1). We also determined the GR expression in cytosolic and mitochondrial extracts of PD1.6TEC– cells that display reduced sensitivity to Dex-induced apoptosis (Fig. 2 C). In contrast with the GC-sensitive parental PD1.6 cells, the GR level was not increased in the mitochondrial fraction of PD1.6TEC– cells treated with Dex (Fig. 2 B, subpanels C and D, lane 4 vs. lane 3). These data indicate that selection of PD1.6 cells toward apoptotic resistance in response to Dex is accompanied with reduced mitochondrial translocation of GR.

    Figure 2. (A) Intracellular staining of GR. PD1.6 cells were treated with 100 nM Dex for 2 h and the mitochondria were stained with 50 nM mitotracker. Rehydrated methanol-fixed cells were incubated with M20 antibodies to GR and FITC-conjugated goat anti–rabbit IgG, and visualized under confocal microscope. Five different cells are presented. (B) Dex induces mitochondrial translocation of GR in PD1.6, but not in PD1.6TEC– cells. PD1.6 cells were incubated in the absence or presence of 100 nM Dex for 2 h before subcellular fractionation using the Oncogene fractionation kit. GR was detected by Western blotting using the PA1-511A antibody to GR (A, C, and D). (C and D) Data were obtained by two different exposure times. The blots were reprobed with antibodies to -tubulin (B) and CytoC (D). CytoC could be used as a mitochondrial marker because the cells were harvested before any CytoC release. (C) Sensitivity of PD1.6 and PD1.6TEC– cells to Dex-induced apoptosis. Cells were either untreated or treated with 100 nM Dex for 20 h, and percentage of apoptotic cells was determined as in Fig. 1 A. (D) The mitochondrial fraction is not contaminated by cytosolic or nuclear proteins. Subcellular fractions were run side by side on the same gel and analyzed by Western blotting using antibodies to -tubulin (A), histone 2B (B), and VDAC (C), which react with cytosolic, nuclear, and mitochondrial fractions, respectively. (E) Dex induces a rapid, temperature-dependent translocation of GR to the nucleus and mitochondria in PD1.6 cells. PD1.6 cells were incubated in the absence or presence of 100 nM Dex for 5 and 15 min at 37°C (lanes 1–3) or 4°C (lanes 4–6) before subcellular fractionation as described in Materials and methods. GR was detected by Western blotting using the PA1-511A antibody to GR (A, C, and E). The blots were reprobed with antibodies to -tubulin (B), histone H2B (D), and VDAC (F). (F) Dex induces sustained GR translocation to the nucleus and the mitochondria in PD1.6 cells. PD1.6 cells were incubated in the absence or presence of 100 nM Dex for 30 min to 5 h and processed as in E. (G) The mitochondrial GR is also immunoreactive to the M20 and P20 antibodies to GR. The mitochondrial samples 1 and 2 of F were rerun on gel and probed with either M20 or P20 antibodies.

    Because the Oncogene kit does not yield nuclear extracts, we applied a cellular fractionation protocol that enabled comparison between mitochondrial and nuclear translocations of GR. Cross-contamination between the cytosolic, nuclear, and mitochondrial fractions was ruled out by using antibodies against -tubulin, histone H2B, and VDAC, respectively (Fig. 2 D). As with the Oncogene kit, this method detected mitochondrial translocation of GR in Dex-treated PD1.6 cells (Fig. 2 E, subpanel E, lanes 2–3 vs. lane 1). A longitudinal follow-up (Fig. 2, E and F) revealed mitochondrial translocation of GR as early as 5 min after addition of Dex, long before the onset of apoptosis. Thereafter, the mitochondrial GR level did not increase further with time (Fig. 2 E and F ). Also, the amount of nuclear GR already peaked at 5 min (Fig. 2, E and F, subpanel C). Similar results were obtained with PD1.6 cells treated with the naturally occurring GC corticosterone (unpublished data). The mitochondrial GR could also be detected on Western blot using the NH2-terminal (aa 5–20)-reacting M20 and the COOH-terminal (aa 750–769)-reacting P20 antibodies (Fig. 2 G), indicating that this GR is in full-length. To exclude the possibility of nonspecific GR binding to the mitochondria, we studied whether this translocation is temperature sensitive. To this end, PD1.6 cells were exposed to Dex at 4°C for 5–15 min before subcellular fractionation. GR did not translocate to either the mitochondria or the nucleus at 4°C (Fig. 2 E, compare lanes 5 and 6 with lanes 2 and 3), indicating that the GR mitochondrial localization is specific and temperature dependent.

    Next, we compared GR translocation to the mitochondria in GC-sensitive PD1.6 cells and GC-insensitive B10 cells (Fig. 3 A). PD1.6 and B10 cells are both derived from a thymic lymphoma, and express similar GR levels. The cells were exposed to 5 and 100 nM Dex for 4 h before subcellular fractionation. Dex induced GR translocation to the nucleus in both PD1.6 and B10 cells (Fig. 3 A, subpanel C, lanes 2, 3, 5, and 6). A dose-dependent mitochondrial translocation of GR was observed in PD1.6 cells (Fig. 3 A, subpanel E, lanes 2 and 3), but not in B10 cells (Fig. 3 A, subpanel E, lanes 5 and 6). In some experiments, a slight increase in GR expression was seen in mitochondrial extracts of B10 cells treated with 100 nM Dex (unpublished data), which reflects residual sensitivity (10–12%) of these cells to Dex (Fig. 1 B). Moreover, Dex induced GR translocation to both the nucleus and the mitochondria in GC-sensitive 2B4 cells (Fig. 3 B) and thymocytes (Fig. 3 C). In 2B4 cells, nuclear translocation of GR was induced by as low as 1 nM Dex, a nontoxic concentration (Fig. 3 B, subpanel C, lane 2). This Dex concentration was insufficient to cause mitochondrial GR translocation in the same cells (Fig. 3 B, subpanel E, lane 2). However, at toxic Dex concentrations (10 and 100 nM), GR translocated to the mitochondria as well (Fig. 3 B, subpanel E, lanes 3 and 4 vs. lane 1).

    Figure 3. (A–E) Dex induces GR translocation to the mitochondria in GC-sensitive PD1.6 cells (A), 2B4 cells (B), and thymocytes (C), but not in GC-resistant B10 (A), NB4 (D), S49 (E), and Jurkat (E) cells. Cells were incubated in the absence or presence of various Dex concentrations for 4 h before subcellular fractionation and Western blotting. In addition, NB4 cells were treated with 1 μM As2O3 (D, lanes 5 and 6), which is known to stabilize PML in these cells (reference 26).

    Contrary to GC-sensitive cells, GC-resistant NB4 (Fig. 3 D, lanes 3–6) and S49 (Fig. 3 E, lanes 1–3) cells responded to Dex with GR translocation to the nucleus, but not to the mitochondria. We also treated NB4 cells with As2O3, which stabilizes PML and degrades the PML–RAR fusion protein (26). As2O3 stabilized GR in these cells (Fig. 3 D, subpanel A, lane 5 vs. lane 3) to a level comparable with that of PD1.6. Even after As2O3 treatment, Dex did not cause mitochondrial translocation of GR in NB4 cells (Fig. 3 D, subpanel E, lane 6). Although S49 cells express a high basal level of GR (Fig. 3 E, subpanel A, lane 1), no GR translocation to the mitochondria is seen (Fig. 3 E, subpanel E, lanes 2 and 3). Thus, mitochondrial GR translocation does not correlate with GR expression but rather with the apoptotic sensitivity to GC.

    Jurkat cells express a very low level of GR (Fig. 3 E, subpanel A, lane 4), which may explain their unresponsiveness to the apoptotic effects of GC. It is well documented that GC-induced apoptosis requires a threshold level of GR (24, 27). Although GR translocates to the nucleus in Dex-treated Jurkat cells (Fig. 3 E, subpanel C, lanes 5 and 6), GR is barely seen in the mitochondria (Fig. 3 E, subpanel E, lanes 5 and 6). Moreover, Jurkat cells express one WT and one mutated (R477H) GR allele, as do the GC-sensitive CCRF–CEM cells (28).

    It should be noted that in most of the GR-expressing cell lines tested, a certain basal level of GR is detected in the mitochondria (e.g., Fig. 3 A , B , and E ). Altogether, the data demonstrate correlation between GR translocation to the mitochondria and sensitivity to GC-induced apoptosis.

    Mitochondrial translocation of GR can occur without concomitant apoptosis

    The GR antagonist RU486 induces a different conformational change of GR than GC agonists (29). It causes nuclear translocation of GR, but prevents its binding to GREs, thereby avoiding transactivation. We sought to find out whether RU486 affects mitochondrial translocation of GR. Initially, we analyzed the effect of RU486 on Dex-induced apoptosis of PD1.6 cells. As expected, RU486 prevented this apoptotic response (Fig. 4 A). When looking on the intracellular trafficking of GR, we found that RU486 induced both nuclear and mitochondrial translocation of GR in PD1.6 cells (Fig. 4 B, subpanel C and E, respectively, lane 3). A combination of RU486 and Dex had an additive effect on GR translocation to the mitochondria (Fig. 4 B, subpanel E, lane 4). These data indicate that a mere localization of GR to the mitochondria is insufficient for inducing apoptosis, which apparently requires a particular GR conformation.

    Figure 4. (A) RU-486 prevents Dex-induced apoptosis of PD1.6 cells. PD1.6 cells were incubated with various concentrations of Dex in the absence or presence of 5 μM RU-486 for 20 h. Percentage of apoptosis was determined as in Fig. 1 A. (B) RU-486 induces mitochondrial translocation of GR in PD1.6 cells. Cells were incubated in the absence or presence of 5 μM RU-486 and/or 100 nM Dex for 2 h followed by subcellular fractionation and Western blotting.

    TEC induces mitochondrial, but not nuclear, translocation of GR in PD1.6 cells

    We have recently shown that thymic epithelial cells (TECs) induce apoptosis of PD1.6 cells in a GR-dependent manner (30). It was therefore of interest to study the intracellular trafficking of GR in PD1.6 cells cocultured with TECs. For this purpose, PD1.6 cells were incubated alone or on a TEC monolayer for 4 h. In PD1.6 cells grown alone, most of the GR is localized in the cytosol (Fig. 5 A, lane 1). After cocultivation with TECs, GR is found in the mitochondrial fraction, but, surprisingly, not in the nuclear fraction (Fig. 5, compare lane 2 in subpanel E vs. subpanel C). PD1.6 cells treated with 100 nM Dex for 4 h were used as a positive control for nuclear translocation. As expected, this treatment caused both nuclear and mitochondrial translocation of GR (Fig. 5, lane 7 in subpanels C and E, respectively). Cocultivation on TEC did not cause a reduction in the cytosolic GR level as seen with Dex (Fig. 5 A, compare lanes 2 and 7 with lane 1). It should be noted that a similar quantity of GR was observed in the mitochondrial fractions of PD1.6 cells cocultivated with TECs and of PD1.6 cells treated with 100 nM Dex (Fig. 5 E, compare lanes 2 and 7). This outcome is consistent with the similar extent of apoptosis induced by these two stimuli (30). To exclude the possibility that GR detected in PD1.6 cells cocultured with TECs is due to TEC contamination, we included in the experiment Dex-resistant PD1.6Dex– cells that express minute amount of GR (30). No GR could be seen in the mitochondrial or nuclear fractions of these cells after cocultivation with TECs (Fig. 5, lane 4 in subpanels E and C, respectively). As a negative control, we used TEC-resistant PD1.6TEC– cells (30), which show only a slight reduction in GR expression level in comparison to the parental PD1.6 cells. Interestingly, TECs induced neither mitochondrial nor nuclear translocation of GR in PD1.6TEC– cells (Fig. 5, lane 6 in subpanels E and C, respectively). Altogether, our data demonstrate that TEC induces GR translocation to the mitochondria in TEC-sensitive PD1.6 cells, but not in TEC-resistant PD1.6TEC– cells. Because TEC causes apoptosis of PD1.6 cells in a GR-dependent manner (30) and induces mitochondrial (but not nuclear) GR translocation, we propose that GR translocated to the mitochondria is a trigger of apoptosis. Our findings also show that the nuclear and mitochondrial translocations of GR are differentially regulated.

    Figure 5. TEC induces mitochondrial, but not nuclear, translocation of GR in TEC-sensitive PD1.6 cells. PD1.6, PD1.6TEC–, and GR-deficient PD1.6Dex– cells were incubated alone or with TEC for 4 h. Nonadherent lymphoma cells were harvested and subjected to subcellular fractionation and Western blotting. The samples were harvested at an early stage after cocultivation with TEC, long before the onset of the apoptotic process. Thus, CytoC could be used as a marker for the mitochondria.

    The mitochondrial localization signal (MLS) and the nuclear localization signal (NLS) are located at different domains of GR

    Our data showing that, depending on circumstances, GR translocates either to the nucleus or to the mitochondria, suggest that nuclear and mitochondrial GR translocations are differentially regulated. We therefore searched for signals directing the intracellular trafficking of GR. To this end, we transfected GR-negative 293 cells with WT or various deletion mutants of human GR (hGR). In the absence of ligand, the transfected GR is expressed in cytosol, nucleus, and mitochondria (Fig. 6 A, lane 1). Addition of Dex did not alter the intracellular distribution of GR in these cells (unpublished data). This pattern of exogenous GR expression enabled the search for NLS and MLS. Using 293 cells transfected with GR deletion mutants, we found that NH2-terminal GR fragments (1–488, 1–515, and 1–550) translocated to the nucleus, but not to the mitochondria (Fig. 6 A, lanes 2–4), indicating that these mutants contain NLS but not MLS. GR428-490 (DBD) and GR490-515 translocated to the mitochondria, but barely to the nucleus (Fig. 6 A, lanes 5 and 6), indicating that NLS resides within aa 428–515 of hGR, which is in accordance with the three NLS described previously (31). GR550-600 translocated to the nucleus, but barely to the mitochondria (Fig. 6 A, lane 7), indicating that MLS resides, at least in part, within aa 550–600. A residual mitochondrial translocation observed with GR550-600 suggests that the 550–600 domain may act in concert with another site within the COOH-terminal region of GR. Similar to WT GR, the 727-777 mutant distributed to both the nucleus and the mitochondria (Fig. 6 A, lane 8). Also, the GR77-262 deletion mutant translocated to both the mitochondria and nucleus (unpublished data). Hence, NLS and MLS are located within different domains of the GR protein.

    Figure 6. (A) MLS and NLS are located within different domains of GR. GR-negative 293 epithelial cells were transfected with plasmids encoding the indicated human GR variants. After 20 h, the cells were subjected to subcellular fractionation and GR was detected on Western blot using the PA1-511 antibody to GR. (B) -Helix wheel model of the putative MLS located within aa 558–580 of human GR. The positive-charged arginine and lysine (red) and the hydrophilic threonine (orange) are located on the one side of the -helix, whereas the hydrophobic aa leucine, isoleucine, valine, and tryptophane (blue) are located on the opposite side of the -helix. Amino acids interacting with GC are labeled with gray numbers. This putative MLS is the -Helix 3 of LBD. (C) The -Helix 3 (aa 558–580) as it appears in the 1M2Z crystal structure. The same color labeling of aa is used as in B. (D) The MLS of GR resembles that of cytochrome C oxidase (COX). Amino acid sequence alignment between MLS of GR and MLS of COX. (E) The R564G and R575G mutants show reduced ability to enter the mitochondria. Mouse GFP-GR was point-mutated at aa 564 or 575 corresponding to the human R558 and R569, and the ability of these mutants to enter the mitochondria was determined as described in A. (F) The MLS (H3) of GR in the LBD crystal structure. The putative MLS is labeled in red in the 1M2Z crystal structure of a dimer complex of the human GR LBD (aa 521–777) bound to Dex and a Tif2 coactivator motif (reference 6).

    The 550–600 domain of GR comprises the -Helix 3 (aa 558–580) of the LBD. This -helix is characterized by a series of positively charged aa (arginine/lysine), hydrophilic aa (threonine), and hydrophobic aa (valine, isoleucine, and leucine), but lacks negatively charged aa. The positively charged and hydrophilic aa locate on the one side of the -helix, whereas the hydrophobic aa locate on its other side (Fig. 6, B and C). This arrangement is compatible with the requirements for an MLS (32). It shows some sequence similarity to the MLS of cytochrome C oxidase (COX) (Fig. 6 D). To verify that the MLS resides within aa 558–580 stretch, Arg564 and Arg575 of mouse GFP-GR (which correspond to human GRArg558 and Arg569, respectively) were each mutated to glycine. Arg575 was also mutated to aspartate. The ability of these mutants to translocate to the mitochondria was compared with that of GFP-GR WT. R564G and R575G showed reduced ability to enter the mitochondria, whereas R575D behaved as WT in this respect (Fig. 6 E). These data indicate that the secondary structure created by R564 and R575, rather than their charge, is important for the MLS integrity. When this MLS is placed in the crystal structure of dimerized ligand-bound LBD, the two MLS appear adjacent (Fig. 6 F). The differential location of MLS and NLS may explain the dissociated nuclear and mitochondrial translocations of GR observed in various cell types in response to diverse stimuli.

    GR targeted to the mitochondria induces apoptosis

    Our data suggest a role for mitochondrial GR in mediating apoptosis. To verify this hypothesis, we sought to distinguish between the effects exerted by GR in the mitochondria and in other intracellular compartments. To this end, we constructed a GR variant (MLSCOX-GFP-GR) that exclusively localizes to the mitochondria. This variant was attained by adding the MLS of COX upstream to GFP-GR. We initially analyzed the intracellular localization of MLSCOX-GFP-GR in the absence or presence of Dex. HeLa, H1299, and PC3 cells transfected with GFP-GR or MLSCOX-GFP-GR were either untreated or treated with 100 nM Dex for 2 h. Thereafter, the cells were stained with red mitotracker to visualize the mitochondria and analyzed by confocal microscopy. The MLSCOX-GFP-GR exclusively localized to the mitochondria both in the absence and in the presence of Dex (Fig. 7, A and B). Hence, this construct was indeed useful for studying the effect of mitochondrial GR on apoptosis. For this purpose, we transfected HeLa cervical carcinoma, PC-3 prostate adenocarcinoma, and L929 E8.2 A3 fibroblast-like cells with plasmids encoding either GFP-GR or MLSCOX-GFP-GR. After 48 h, the percentage of apoptotic transfectants was compared with that of nontransfectants from the same sample. GFP-GR induced apoptosis of HeLa and L929 E8.2 A3 cells, but not of PC-3 cells (Fig. 7 C). MLSCOX-GFP-GR induced apoptosis of HeLa and L929 E8.2 A3 cells more efficiently than GFP-GR (Fig. 7 C). Interestingly, PC-3 cells, which do not undergo apoptosis by GFP-GR, were sensitive to MLSCOX-GFP-GR (Fig. 7 C). These results demonstrate that when GR is directed to the mitochondria, it is capable of inducing apoptosis.

    Figure 7. (A and B) MLSCOX-GFP-GR is exclusively localized to the mitochondria. The mitochondria-directed GFP-GR variant was prepared by inserting the MLS of COX upstream to GFP-GR. HeLa cells (A), H1299 (B), or PC-3 (B) cells were transfected with either GFP-GR or MLSCOX-GFP-GR and stained with red mitrotracker to visualize the mitochondria before confocal microscopy. (C) MLSCOX-GFP-GR is more efficient in inducing apoptosis than GFP-GR. HeLa, L929 E8.2 A3, and PC-3 cells were transfected with plasmids encoding MLSCOX-GFP-GR or GFP-GR. After 48 h, the percentage of apoptotic GFP-positive cells (transfectants) was compared with that of GFP-negative cells (nontransfectants) of the same sample. (D) NLS-defective GRK513-515A induces apoptosis of HeLa cells. Cells were transfected with plasmids encoding GFP-GRwt, GFP-GRK513-515A, or pEGFP-F. After 48 h, the percentage of apoptotic cells was determined as in C. (E and F) A nucleus-directed GFP-GR variant (NLSTAg-GFP-GR) induces apoptosis of HeLa (E) and H1299 (F) cells. Cells were transfected with plasmids encoding GFP-GR, MLSCOX-GFP-GR, NLSTAg-GFP-GR, or pEGFP-F. After 48 h, the percentage of apoptotic cells was assessed as in C.

    NLS-defective GR and a nuclear-directed GR possess proapoptotic properties

    The aforementioned data indicate that mitochondrial GR triggers apoptosis induced by GC. We cannot, however, exclude the possibility that GR may act at additional intracellular sites. To further study in which intracellular compartments GR may induce apoptosis, we analyzed the apoptotic ability of an NLS-defective GR mutant (pEGFP-ratGRK513-515A) and a nucleus-only directed GR variant (NLSTAg-GFP-GR). The latter was constructed by adding the NLS of SV40 T antigen in triplet upstream to GFP-GR of mouse origin.

    The apoptotic ability of pEGFP-GRK513-515A was compared with that of the parental pEGFP-GR WT in HeLa cells. It should be noted that pEGFP-GRK513-515A is unable to translocate to the nucleus (31). We observed that both GR WT and the NLS-defective GR variant induced apoptosis of HeLa cells (Fig. 7 D). pEGFP-F, used as a negative control, did not cause apoptosis of HeLa cells (Fig. 7 D). The ability of pEGFP-GRK513-515A to trigger apoptosis demonstrates that GR may mediate apoptosis by a nucleus-independent manner.

    Last, we compared the apoptotic ability of the nucleus-directed NLSTAg-GFP-GR to those of MLSCOX-GFP-GR and parental GFP-GR. For this purpose, HeLa and H1299 cells were transfected with the respective plasmids. Both NLSTAg-GFP-GR and MLSCOX-GFP-GR were more efficient than GFP-GR in inducing apoptosis of these cells (Fig. 7, E and F). pEGFP-F did not induce apoptosis under the same circumstances (Fig. 7, E and F). These results suggest that an overexpressed GR may induce apoptosis when present either in the mitochondria or in the nucleus. Under physiological conditions, we anticipate that the mitochondrial GR cooperates with nuclear GR in inducing apoptosis.

    DISCUSSION

    Numerous studies have been performed to elucidate the mechanisms by which GC induces apoptosis (2, 3, 5, 27). Several biochemical changes occurring immediately after exposure to GC have been characterized. These include Ca2+ mobilization, activation of Src and Cdk2 kinases, and activation of phosphatidylinositol-specific phospholipase C and acidic sphingomyelinase with subsequent ceramide generation (3, 33–35). Downstream effector mechanisms of GC-induced apoptosis have also been defined. These involve the mitochondria apoptotic pathway mediated by Bax, Bak, Bim, and tBid, and antagonized by Bcl-2 and Bcl-XL (3, 36). Dissipation of the mitochondrial membrane potential (m) is followed by release of cytochrome C and Smac/Diablo to the cytosol (3, 36), which in turn leads to the activation of caspase-9, caspase-3, and endonucleases (3, 36). With the good knowledge of downstream effectors in GC-induced apoptosis, little is known about the role and fate of GR in this response, as well as the reasons why some GR-expressing cells are sensitive, whereas others are not.

    GR expression above a threshold level is necessary, but is not sufficient to activate GC-mediated apoptosis (2, 5, 27). Several studies have indicated that GC-resistant cells may express similar GR levels as GC-sensitive cells (2, 37–39). Likewise, in the present study, we show that GC-sensitive (PD1.6, 2B4, thymocytes) and some GC-resistant (B10, S49) cells express high GR levels. These observations suggest that the apoptotic response is regulated by additional factors acting downstream to ligand–receptor interaction. One possibility is that only a certain isotype of GR can initiate apoptosis. Indeed, various mRNA transcripts of GR have been observed in both mouse and human cells (19, 40, 41); among them, the 1A variant is most abundantly expressed in hematopoietic cells (19, 20). The 1A transcript is mainly translated to a 90-kD GR and a small proportion of a higher molecular mass GR (150 kD) (18). The various cells analyzed in our study mainly express the 90-kD form corresponding to GR. We also detected a small amount of 150-kD GR in the cytosolic fraction of PD1.6, 2B4, B10, and S49 cells (unpublished data). Hence, the 150-kD GR form is not expressed exclusively in cells sensitive to GC, but in GC-resistant cells as well.

    Another factor that could affect GC-mediated apoptosis is the intracellular trafficking of GR. GC causes nuclear translocation of GR in both GC-sensitive and GC-resistant cells. Both transactivation-deficient and transrepression-deficient GR variants have been shown to restore GC sensitivity in human T-ALL cells (7, 14, 42, 43). These data together with our present finding that an NLS-deficient GR mutant is proapoptotic suggest that nuclear-independent mechanisms are likely involved in this death pathway.

    It has been suggested that mGR is involved in GC-induced apoptosis based on the finding that a cDNA derived from full-length 1A transcript, imparted both mGR expression and GC sensitivity to some GC-resistant cells (18). However, this cDNA also caused GR overexpression in the cytosol (18), which may have contributed to the GC sensitivity of the cells. In the present paper, we have further studied the role of mGR in GC-induced apoptosis by analyzing mGR expression in GC-sensitive and resistant lymphoid cells. mGR was expressed on some cell types that were GC resistant, but not on the GC-sensitive cells studied. Thus, mGR is not required for GC-induced apoptosis, and its mere presence does not impose susceptibility to GC. Our findings are compatible with those of Gametchu et al. (17), showing that mGR-negative lymphoma cells may be GC sensitive.

    Another possibility could be that mitochondrial GR is the trigger of apoptosis. GR has been shown to be located within the mitochondria in some cell types (22–24). In these studies, GR was detected in the mitochondrial membranes and in the matrix space. Likewise, we also observed a basal mitochondrial GR expression in most of the lymphoma cells analyzed. However, Dex induces GR translocation to the mitochondria in GC-sensitive, but not in GC-resistant, cells. This is in contrast with nuclear translocation of GR that takes place in all cell types. This is the first qualitative difference in GR behavior described that distinguishes between GC-sensitive and resistant cells, suggesting a role for mitochondrial GR in apoptosis.

    In GC-sensitive cells, mitochondrial and nuclear translocations of GR occur simultaneously within the first minutes after exposure to Dex. After its initial translocation, the amount of GR in the mitochondria was maintained at a steady state. Thus, the elevated mitochondrial GR level is sustained in GC-sensitive cells, which is in contrast with the transient residence of GR in the mitochondria (5–30 min) in HeLa cells and with the reduced amount of mitochondrial GR in a glioma cell line after Dex treatment (24, 44). The sustained expression of GR in the mitochondria of GC-sensitive cells may account for the apoptotic effects of GC. Another support for a role of mitochondrial GR in mediating apoptosis comes from the observation that TECs, which trigger apoptosis of PD1.6 in a GR-dependent manner (30), induces GR translocation to the mitochondria, but not to the nucleus. Thus, GR translocation to the nucleus is not necessary for the apoptotic response. However, we cannot exclude the possibility that mitochondrial GR cooperates with nuclear GR in inducing apoptosis. The fact that a mitochondria-directed GR induces apoptosis suggests that exclusive expression of GR in the mitochondria is sufficient for triggering apoptosis. Conversely, a nucleus-directed GR is also proapoptotic. It should be noted that the latter data were obtained by a transient transfection assay where the proteins were overexpressed. Thus, the GR effects are more pronounced than at physiological GR levels. Nevertheless, it is a valuable approach that provides information on GR function. Altogether, our data show that mitochondrial translocation of GR correlates with apoptotic sensitivity.

    This behavior of GR resembles that of the proapoptotic p53 and Nur77 proteins. These proteins have recently been shown to have a direct apoptogenic role at the mitochondria, besides their nuclear effects (45, 46). Our results add GR to the growing list of proteins that mediate apoptosis when localized to the mitochondria. Also, the thyroid, estrogen ? and retinoid X receptors have been shown to be located in the mitochondria (24, 47, 48). Thus far, however, the mitochondrial localization of these receptors has not been implicated in apoptosis.

    The mechanism by which mitochondrial GR mediates apoptosis is a matter for further study. GC has been shown to regulate mitochondrial transcription and energy production (24), and GRE elements have been found in the mitochondrial genome (49). It should be mentioned in this regard that some of the apoptotic effects of p53 and Nur77 occur independently of their transcriptional activities (45, 46). p53 and Nur77 interact with the protective Bcl-XL and Bcl-2 proteins in the mitochondria (46, 50). It would, therefore, be interesting to find out whether mitochondrial GR has similar effects.

    Another important finding of our study is that nuclear and mitochondrial GR translocations are differentially regulated. For instance, Dex induces both mitochondrial and nuclear GR translocations in GC-sensitive lymphoid cells, but only nuclear translocation in GC-resistant cells. In contrast, TEC induces mitochondrial, but not nuclear, GR translocation. Furthermore, we have partially characterized a putative MLS comprising the -Helix 3 (aa 558–580) of LBD. This domain lies COOH terminally to the three NLSs (aa 428–515) (31), and possesses several traits of an MLS (32). The hydrophobic and positively charged residues are partitioned on opposite sides of the helix. Most of the proteins destined to the mitochondria contain an NH2-terminal mitochondrial transfer peptide that is cleaved off after entering the mitochondrial matrix (32). Some mitochondrial proteins, however, have a noncleavable internal MLS (32). The GR detected by us in the mitochondria is of a similar size as cytosolic GR, indicating that it does not undergo cleavage upon mitochondrial translocation. Hence, GR contains a noncleavable internal mitochondrial targeting sequence possessing an amphipathic presequence-type helix. This target sequence resembles the internal MLS of BCS1 (51).

    It is conceivable that GR is transported to the mitochondria by a heat-shock protein, as its 558–580 domain overlaps with one of several characterized Hsp90-binding sites (5), and both Hsp90 and Hsp70 function as chaperones that interact with the mitochondrial protein import receptor Tom 70 (52). Further studies are required to verify the role of heat shock proteins in GR trafficking.

    In summary, we conclude that mitochondrial GR acts independently of nuclear GR in inducing apoptosis, and that mitochondrial and nuclear GR translocations are differentially regulated. Given the multiple nuclear adverse effects of GC therapy, our findings propose further research focusing on the development of therapeutic modalities that preferentially direct GR to the mitochondria.

    MATERIALS AND METHODS

    Cells.

    PD1.6 thymic lymphoma (30), B10 thymic lymphoma (30), S49 T lymphoma (provided by J. Hochman, The Hebrew University of Jerusalem, Jerusalem, Israel), TEC (provided by A. Kruisbeek, The Netherlands Cancer Institute, Amsterdam, Netherlands), 293 kidney epithelial cells, HeLa cervical carcinoma, and L929 E8.2 A3 fibroblast-like cells (provided by W.V. Vedeckis, Louisiana State University, New Orleans, LA) were grown in DMEM supplemented with 10% heat-inactivated FCS, 2 mM glutamine, 10 mM Hepes, 1 mM sodium pyruvate, nonessential aa, antibiotics, and 50 μM ?-mercaptoethanol. 2B4 T cell hybridoma, 4B2 PML, Jurkat ALL, thymocytes, H1299 lung adenocarcinoma, and PC-3 prostate adenocarcinoma were cultured in RPMI 1640 with the same supplements as for DMEM. PD1.6Dex– cells were derived from PD1.6 cells by repeated exposure to increasing concentrations of Dex and PD1.6TEC– cells were derived from PD1.6 cells repeatedly cocultured with TECs (30).

    Plasmids.

    The following plasmids were used: GFP-mouse GR (provided by L.J. Muglia, Washington University in St. Louis, St. Louis, MO); hGRDBD and hGR77-262 (provided by W. Doppler, Universit?t Innsbruck, Innsbruck, Austria); hGR WT, hGR1-488, hGR1-515, hGR1-550, hGR418-777, hGR490-515, hGR550-600, hGR550-777, and hGR727-777 (provided by T.D. Gelehrter, University of Michigan, Ann Arbor, MI); pEGFP-rat GR and pEGFP-ratGR K513-515A (provided by K.R. Yamamoto, University of California, San Francisco, CA); pCMV/myc/mito (Invitrogen) (provided by S. Ostrand-Rosenberg, University of Maryland, Baltimore, MD); pCMV/myc/nuc (Invitrogen) (provided by D. Bensi, Mayo Clinic, Rochester, MN); farnesylated GFP (pEGFP-F; CLONTECH) and pEGFP-N1 (CLONTECH) (provided by Y. Haupt, The Hebrew University of Jerusalem, Jerusalem, Israel). A mitochondria-directed GFP-GR variant (MLSCOX-GFP-GR) was prepared by inserting the MLS of COX (MSVLTPLLLRGLTGSARRLPVPRAKIHSL) NH2-terminal to GFP-GR. A PCR product containing this MLS was obtained from pCMV/myc/mito using 5'-primer CTAGCTAGCTGACGCAAATGGGCGGTAGGCGTG-3' harboring a NheI site and 3'-primer CCCACCGGTTTGGCCCCATTCAGATCCTCTTC-5' harboring a AgeI site. After double digestion with NheI and AgeI, the PCR product was inserted within the NheI/AgeI sites of GFP-GR. A nucleus-directed GFP-GR variant (NLSTAg-GFP-GR) was prepared by inserting the NLS of SV40 T antigen in triplet () NH2-terminal to GFP-GR. pCMV/myc/nuc was used as template for the aforementioned primers. Mutations in GR were introduced by site-directed mutagenesis (QuickChange kit; Stratagene). Sequencing of plasmids was done at the DNA Sequencing Facility of The Hebrew University of Jerusalem, Israel.

    Indirect immunofluorescence.

    Cell surface staining of GR was performed by incubating cells with M20 antibody to GR (Santa Cruz Biotechnology) followed by FITC-conjugated AffiniPure F(ab)2-fragment of goat anti–rabbit IgG (Jackson ImmunoResearch Laboratories). Intracellular GR staining was performed on rehydrated methanol-fixed cells.

    Determination of apoptosis.

    Apoptosis was assessed by cell cycle analysis and caspase 3 activation. For cell cycle distribution, cells were fixed with ice-cold methanol, rehydrated in PBS, and treated with 50 μg/ml RNase. After adding 5 μg/ml propidium iodide, the DNA content was analyzed by flow cytometry (FACSCalibur; Becton Dickinson). Subdiploid cells are regarded as apoptotic cells, and presented as percentage of the whole cell population. Caspase 3 activation was analyzed by incubating rehydrated methanol-fixed cells with antibody to cleaved caspase 3 (Asp 175; Cell Signaling Technologies) followed by FITC-conjugated AffiniPure F(ab)2-fragment of goat anti–rabbit IgG (Jackson ImmunoResearch Laboratories).

    Subcellular fractionation and Western blot.

    Cell pellets were gently resuspended in cytoplasmic buffer (10 mM Hepes, pH 7.4; 1.5 mM MgCl2, 10 mM KCl, 0.5 mM DTT, 10 mM Na2MoO4, 2 mM PMSF, 20 μg/ml aprotinin, 0.1% NP-40; 25 mM NaF, and 0.2 mM Na3VO4) and kept on ice for 10 min before centrifugation at 900 g for 10 min. The nuclear pellets were processed as described below. The cytoplasmic supernatant was recentrifuged at 900 g to ensure complete removal of nuclear material. The resulting supernatant was centrifuged at 10,000 g for 30 min. The cytosolic supernatant was processed for Western blot by adding protein sample buffer (PSB) x 4.5. The mitochondrial pellet was washed with cytoplasmic buffer, recentrifuged at 10,000 g, and dissolved in PSB x 1.5. The nuclear pellet was washed with cytoplasmic buffer and recentrifuged at 900 g before their extraction in nuclear buffer (20 mM Hepes, pH 7.4, 1.5 mM MgCl2, 420 mM NaCl, 25% glycerol (vol/vol), 0.2 mM EDTA, 0.5 mM DTT, 10 mM Na2MoO4, 2 mM PMSF, 20 μg/ml aprotinin, 25 mM NaF, and 0.2 mM Na3VO4). The nuclear extracts were cleared at 20,000 g for 10 min, and processed for Western blot by adding PSB x 4.5. GR was detected on Western blot by using the PA1-511A (Affinity BioReagent), M20, or P20 antibodies to GR (Santa Cruz Biotechnology). The blots were reprobed with antibodies to -tubulin (Sigma-Aldrich), cytochrome C (CytoC; Santa Cruz Biotechnology), VDAC (Oncogene Research Products), or histone H2B (LG2-2; provided by M. Monestier, Temple University, Philadelphia, PA) (53). The Oncogene cytosol/mitochondria fractionation kit (QIA88; Oncogene Research Products) was used according to the manufacturer's instructions.

    Cocultivation of PD1.6 cells with TECs.

    5 x 106 TECs were seeded in 75 cm2 tissue culture bottles (Nunc). On the following day, 107 PD1.6 cells were added to the TEC monolayer in a 40-ml medium. After 4 h, nonadherent PD1.6 cells were harvested and processed for cell fractionation as described before.

    Transient transfection assay.

    Cells were transfected with the given plasmids by using the calcium phosphate precipitation method. For apoptosis assay, the adherent cells were trypsinized and collected in culture supernatants. Flow cytometry was performed while gating on GFP positive or negative cells. Percentage of apoptotic transfectants (GFP-positive) was compared with that of nontransfectants (GFP-negative) from the same culture. For confocal microscopy, the cells were incubated with red mitotracker (50 nM; Invitrogen) during the last 30 min of incubation. For cell fractionation studies, adherent transfected cells were harvested with rubber policeman and centrifuged at 720 g for 5 min.

    Acknowledgments

    We thank Dr. M. Tarshish for excellent assistance with confocal microscopy.

    This work was supported by The Concern Foundation, CA.

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