| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
* Laboratory of Biochemistry, Institute for Interfacial Engineering, University of Stuttgart and Fraunhofer Institut für Grenzflächen-und Bioverfahrenstechnik, Stuttgart, Germany;
Gaubius Laboratory, Netherlands Central Organization for Applied Scientific Research-Prevention and Health, Leiden, The Netherlands; and
Department of Biochemistry and Molecular Cell Biology, Institute of Biochemistry, University Hospital Rheinisch-Westfälische Technische Hochschule, Aachen, Germany
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
The mechanisms of action of MIF have not yet been elucidated. In particular, a membrane receptor for MIF has not yet been identified. However, MIF function may be mediated, at least in part, by other molecular pathways. MIF has been demonstrated to exhibit two distinct enzymatic activities, i.e., a Pro2-dependent tautomerase and a Cys57-Ala58-Leu59-Cys60 (CALC)-dependent oxidoreductase activity (13, 14, 15). The enzymatic activities of MIF could represent intracellular functions of this mediator. It has been suggested that the catalytic redox activity of MIF plays a role in the regulation of cellular redox stress (16). In this respect, MIF is reminiscent of another immune mediator, thioredoxin (Trx) or adult T cell leukemia-derived factor, that also, through its enzymatic thiol-protein oxidoreductase activity, participates in the regulation of cellular redox stress (17, 18). Although a link between the catalytic properties of MIF and its immunoregulatory activities has not yet been unanimously derived (19, 20), it is likely that MIF-mediated "enzymatic signaling" may critically contribute to at least some of the effects of MIF, as, for example, the enzyme-dead mutant C60SMIF does not exhibit certain MIF-like macrophage-activating properties (15, 21). Furthermore, a physiological involvement of MIF in the regulation of the cellular redox state is supported by studies showing that MIF binds to and regulates peroxiredoxin (PAG) (22) and that MIF functions as a redox regulator in the ischemic myocardium (23).
Another clue toward the elucidation of the molecular pathways of MIF action has come from the identification of the coactivator Jun activation domain binding protein 1 (JAB1) as an intracellular binding protein of MIF (24). Following nonreceptor-mediated endocytosis, MIF specifically interacts with JAB1 in both immune and nonimmune cells (24, 25). Intracellular interaction of MIF and JAB1 also occurs in MIF-producing cells (24). MIF can modulate AP-1 transcriptional and c-Jun N-terminal kinase (JNK) pathways and the cell cycle through JAB1. The CALC redox motif of MIF is important for at least some of the JAB1-mediated activities of MIF (24). Other studies have suggested that MIF can also affect the extracellular signal-regulated kinase 1/2 pathway and cyclooxygenase activity as well as tyrosine kinase signaling (26, 27), but it is unknown through which signal-mediating proteins these effects occur.
The cellular effects of MIF reported include effects on cell proliferation and differentiation, cell migration, and on the gene expression of other inflammatory mediators (summarized in Refs. 3 and4). Depending on the physiological context and the cell type, MIF can stimulate or inhibit cell proliferation (3, 24, 28). MIF down-regulates the expression of the cell cycle regulators p21 and cyclin G, while it stabilizes p27 levels (24, 29). MIF inhibits the random migration of monocytes/macrophages but down-regulates the migratory properties of chemokine-stimulated monocytes and lymphocytes and enhances the migration of tumor cells (20, 30, 31). MIF induces the gene expression of inflammatory cytokines such as TNF and IL-1
and, depending on the context, can induce or down-regulate inducible NO synthase and IL-6 (7, 32, 33). Together, this indicates that MIF participates in the regulation of cellular activity in a complex and multiple fashion.
Apoptosis is a key cellular event, contributing, among many other processes, to the degree of immune activation and inflammation under various conditions. Recently, Hudson et al. (29) showed that MIF participates in the regulation of apoptosis. MIF endogenously overexpressed in macrophages could overcome apoptosis induced by the tumor suppressor p53 and MIF inhibited p53-mediated transcriptional activation. Suppression of p53-mediated apoptosis by MIF represents a novel molecular link between inflammation and cancer (29). Regulation of p53 by MIF was confirmed in a very recent study on the role of MIF in the innate immune response (34). Of note, identification of a specific interaction between p53 and JAB1 (35) and MIF and JAB1 (24) could provide a connection between apoptotic processes, the regulation of cellular stress by MIF, and the JAB1- and p53-dependent pathways of MIF action.
In this study, we investigated these complex mechanistic interactions by asking whether MIF was able to participate in the regulation of pro-oxidative stress-induced cellular apoptosis. First, we established experimental systems to more generally establish the antiapoptotic activity of MIF. As MIF had previously been reported to act by both typical cytokine-like, transcellular pathways and by intracellular mechanisms, we probed the effects of both exogenously added biologically active rMIF and endogenous MIF overexpressed specifically through a tetracycline (tet-off; tet) gene regulation cassette. The potential role of the enzymatic redox activity of MIF for its apoptosis-regulating properties was tested by comparing the effects of wild-type (wt) MIF with those of the enzyme-dead mutant C60SMIF. The specific effects of MIF on pro-oxidative stress-induced apoptosis were evaluated by studying apoptosis in a cellular model of mild pro-oxidative stress, through redox stress induction by diamide, and by measuring intracellular glutathione (GSH) levels. The activity of stress-activated protein kinase/JNK was studied in the context of pro-oxidative stress.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
The thiol reagents L-cystine (L-cys), 2-ME, GSH, and DTT as well as camptothecin (cam), doxycycline (dox), and diamide were obtained from Sigma-Aldrich (Deisenhofen, Germany). cam was dissolved in DMSO at a concentration of 2 mg/ml, the stock solution was aliquoted and kept at -20°C for up to 6 mo. The dox stock solution was prepared in bidistilled water at a concentration of 1 mg/ml and stored at 4°C for up to 5 days. Oligonucleotide primers were acquired from Life Technologies (Eggenstein, Germany). The SuperFect transfection reagent was from Qiagen (Hilden, Germany) and the other molecular biology reagents were from Life Technologies or New England Biolabs (Heidelberg, Germany). 2-nitrophenyl--D-galactopyranoside was from Roche Diagnostics (Mannheim, Germany). Thiol-free RPMI 1640 medium was bought as custom-made medium from Life Technologies and BioWhittaker (Verviers, Belgium) to be free of L-cys, L-GSH, and 2-ME. Dialyzed FCS was obtained from Life Technologies. Thiol-free RPMI 1640 medium supplemented with 10% dialyzed FCS was termed thiol-free medium (TFM). Miscellaneous cell culture reagents, antibiotics, and other media were from Life Technologies and all other miscellaneous chemicals and salts were from Sigma-Aldrich. All reagents were of the highest grade commercially available.
Recombinant proteins and constructs
The pET11b plasmid was from Novagen/Calbiochem (Heidelberg, Germany). The cloning, expression, purification, and renaturation of wt human MIF and of the redox mutant C60SMIF have been described (15, 21, 36).
The following tet system-related plasmids were used: pUHD15–1neo, the regulator plasmid for both the transient and stable transfections containing the tet-controlled transcriptional activator gene and a neomycin resistance gene; pUHD10–3, the response plasmid for both the transient and stable transfections containing the target gene cloned after the tet-responsive element and a minimal CMV promoter; pUHD16–1, a response plasmid for monitoring the efficiency of the transient transfections containing a lacZ reporter gene cloned after a constitutive CMV promoter; pX343, a response plasmid containing a hygromycin resistance gene for the selection of the stably transfected clones; pUHG16–3, a response plasmid containing a lacZ gene cloned after the tet-responsive element and a minimal CMV promoter that was used to control for tet-controlled transcriptional activator activity in tet-regulated HeLa cells (HtTA; see below) and for the screening for high switch-on factors of the target gene. These constructs as well as the HtTA cell line were a kind gift from Prof. H. Bujard (Center for Molecular Biology, Heidelberg, Germany). The construction of these tet system reagents has been described (37).
The construction of a human MIF cDNA-bearing tet-responsive plasmid, pMIFRP, for the transient and stable transfection experiments, was performed by PCR cloning. The human MIF sequence was amplified from the pET11b-huMIF plasmid (15) using the SacII restriction site containing the 5' primer: 5'-GCG CCG CGG ATG CCG ATG TTC ATC GTA AAC AC-3' and the BamHI restriction site containing the 3' primer: 5'-CGG GAT CCT TAG GCG AAG GTG GAG TTG TTC CAG C-3'. PCR-amplified DNA was ligated into the pUHD10–3 vector digested with SacII and BamHI. pC60SMIFRP was constructed in an identical manner except that a pET11b plasmid bearing the human C60SMIF cDNA sequence (15) was used as template in the amplification step.
All generated cDNA sequences were confirmed by bidirectional DNA sequencing.
Cell culture
Cell lines were obtained from the Deutsche Sammlung von Mikroorganismen und Zellkulturen (Braunschweig, Germany) or American Type Culture Collection (Manassas, VA) unless stated otherwise. Human Jurkat T cells were a kind gift from Dr. F.-J. Johannes (Fraunhofer Institut für Grenzflächen-und Bioverfahrenstechnik, Stuttgart, Germany). Cells were cultured by routine protocols at 37°C in a humidified incubator with 5% CO2.
HtTA cells represent HeLa cells stably transfected with a tet regulator gene cassette (see above). HtTA cells stably expressing the human MIF gene (HtTAM) were generated by transfecting HtTA cells with pMIFRP using the SuperFect transfection reagent. HtTA cells were transfected with pMIFRP and pX343 that had been linearized with AseI and PvuII, respectively, before transfection. The plasmids were applied at a ratio of pMIFRP/pX343 of 1:10. Cells were incubated overnight, transferred to 10-cm cell culture dishes and incubated for 13 days in the presence of 250 µg/ml hygromycin. From a total of 36 positive clones, 34 were transferred to 24-well plates, expanded, and clones carrying the human MIF sequence were stably integrated in their genome further selected through G418 and hygromycin treatment as described (37). Three clones were finally obtained and analyzed for their responsiveness to dox and for their MIF expression levels. To determine the efficiency of target gene expression and the switch-on factor of the MIF gene, the cell clones were plated in 3.5-cm dishes at a density of 1 x 105 (1 ml of medium), incubated for 72 h in presence vs absence of dox (1 µg/ml), and analyzed by Bradford protein assay and MIF ELISA (see Results and Fig. 1A). Clone 13 was used for the apoptosis studies.
|
For experiments applying the HtTAM cells, 5 x 106 cells were transferred to 75-cm2 flasks and the cells were incubated for 48–72 h in the presence vs absence of 1 µg/ml dox. Cells were lysed in the appropriate lysis buffer and analyzed for -galactosidase activity, MIF concentration, apoptosis, GSH content, or phospho-c-Jun levels (see below).
Transient transfection experiments were performed differently for each cell line used and condition applied (see below).
Transient transfection and apoptosis assays with Kym-1, HL-60, and HtTA cells
Apoptosis assays following transient transfection of Kym-1 cells with the pMIFRP and pC60SMIFRP plasmids were generally performed according to the following protocol: 2–5 x 105 cells were seeded in 3.5-cm dishes (2 ml of culture medium) the day before transfection. At the time of transfection, the cells had reached 
80% confluency. Transfections were performed with 2–3 µg of the respective plasmid according to the SuperFect protocol. Medium was removed by gentle aspiration and the cells were washed with 2 ml of PBS. Afterward, the cells were incubated for 48 h in the presence or absence of 1 µg/ml dox.
For culture stress-induced apoptosis, cells were harvested thereafter and lysed for subsequent analysis. For cam-induced apoptosis studies, fresh cell culture medium, containing serum (full medium; for TFM see below), 1 µg/ml dox or control solution, and 2 µg/ml cam or control solution was added to the washed cells and the cells were incubated for another 24 h before harvesting and analysis.
For thiol starvation stress-induced apoptosis experiments following transient transfection HtTA (clone HtTA1-1), the cells were grown in full medium (FM) as described above. For the transfections, 5 µg of the respective plasmid were added. The lacZ reporter plasmid pUHG 16–3 was used as a control. After the transfection, cells were incubated in FM containing 1 µg/ml dox or control solution for 30 h. Treatment with TFM in the presence vs absence of dox was for another 18 h. Switch-on levels of MIF and C60S in these cells were controlled by Western blotting analysis using 4–12% NuPAGE gels (Invitrogen; for the blotting and staining procedure see below). Equal amounts of lysate protein as determined by Bradford protein assay were loaded in each lane. A polyclonal rabbit anti-MIF Ab (MIF Ka565), which recognizes the wtMIF and mutant C60SMIF proteins with equal efficiency, was used for detection.
For probing the effects of rMIF and rC60SMIF on apoptosis in Kym-1 cells, the cells were plated at a density of 2 x 106 in 3.5-cm culture dishes and rested for 24 h. MIF or C60SMIF were added at the indicated concentrations and the cells were incubated for 48 h. cam or control solution (2 µg/ml) was added and cells were incubated for another 24 h before harvesting and analysis by cell death ELISA (see below).
HL-60 promyeloblasts were used to compare the effects of rMIF and rC60SMIF on apoptosis induced by cam treatment and to study the effect of rMIF on thiol starvation-induced apoptosis. For the cam-induced apoptosis experiments, HL-60 cells were cultured in RPMI 1640 medium supplemented with 10% FCS, 2 mM L-glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin at a density of 2 x 105. The rMIF proteins were added and the cells were incubated for 24 h. cam or control buffer (2 µg/ml) was added and the cells were incubated for another 24 h. For apoptosis experiments in TFM, HL-60 cells were grown in FM as described above. Before treatment, the cells were washed once with PBS and twice with TFM and were incubated in TFM for 6 h. After another wash with TFM, 200 µM L-cystine, 2 µM 2-ME, or rMIF at the indicated concentrations was added and the cells were incubated for 20 h. Apoptosis analysis of HL-60 cells was performed by annexin V/propidium iodide (PI)-based FACS and cell death ELISA (see below).
Apoptosis assays with HtTAM cells
Apoptosis in HtTAM cells was induced by cam, thiol starvation, or diamide treatment. For cam-induced apoptosis experiments, cells were cultured in the presence vs absence of dox (see above) and treated with cam (0.1–1 µg/ml) for 18 h and cell lysates were analyzed by the cell death ELISA. For thiol starvation-induced apoptosis studies in HtTAM cells, cells were cultured in the presence vs absence of dox as described above, washed extensively in TFM (see above), and cultured in TFM for up to 18 h. For diamide treatment, cells were incubated with or without dox for 48 h as above, diamide was added at different concentrations, and cells were incubated further for up to 2 h. Apoptosis was then assessed by a cell death ELISA, PI staining and microscopy, annexin V/PI-based FACS, or by DNA fragmentation (see below).
-Galactosidase assay
Cell lysis was achieved by adding a buffer containing 25 mM Tris-phosphate buffer, pH 7.8, 2 mM DTT, 2 mM bis-1, 2-diaminocyclohexane-N, N, N',N' tetraacetic acid (CDTA), 10% glycerol, and 1% Triton X-100. To determine the -galactosidase activity, 10 µl of each lysate was mixed with 900 µl of substrate solution, containing 100 mM sodium phosphate buffer, pH 7.0, 10 mM potassium chloride, 1 mM magnesium sulfate, 50 mM 2-ME, and 4 mg/ml 2-nitrophenyl--D-galactopyranoside in a 1.5-ml tube. The reaction was conducted for 2 h at 37°C and was terminated by adding 500 µl of a 1 M bicarbonate solution, pH 9.5. The resulting absorbance was measured at 420 nm using the DU-65 UV-VIS spectrophotometer from Beckman (Munich, Germany).
UV stress-induced apoptosis in Jurkat T cells
UV stress-induced apoptosis was studied in Jurkat T cells. Cells were incubated with or without rMIF for 16 h and subsequently treated with UV light for 1 or 6 h using a Stratalinker (Stratagene, Heidelberg, Germany; 3.6 Joule/cm2).
Cell death ELISA
The degree of cellular apoptosis was mainly scored by a commercial apoptosis-specific cell death ELISA from Roche Diagnostics (kit no. 1544675; Mannheim, Germany). The ELISA provides a quantitative determination of histone-associated DNA fragments (mono- and oligonucleosomes) in the cytoplasmic fraction of a cell lysate. The presence of mono- and oligonucleosomes is a typical feature of cells undergoing apoptosis.
Briefly, the assay represents a sandwich ELISA applying mAbs directed against DNA and histones. First, the anti-histone Ab (clone H11-4) was adsorbed to the microtiter plate. Following blocking, the cell lysates containing the free nucleosomes were added. During the incubation step, the nucleosomes, via their histone components (histone H2B), bound to the immobilized anti-histone Ab. Next, the formed complexes were incubated with an anti-DNA Ab conjugated with peroxidase (clone MCA-33). Immune complexes were then determined photometrically using ABTS as a substrate. Quantification was performed by spectrophotometric measurement at 405 nm.
FACS assay and fluorescence microscopy
For FACS analysis of HtTAM cells, cells were cultured in normal medium in the presence vs absence of dox for 48 h. Cells were washed with PBS and with TFM medium. dox or control solution (1 µg/ml) was added and the cells were cultured in TFM for another 18 h. HL-60 cells were rested in normal medium and then incubated with TFM for 18 h. Afterward, the cell samples were analyzed with the annexin V-FLUOS staining kit (Roche Diagnostics) according to the manufacturer’s instructions. Briefly, medium was removed from the cells and the cells were washed in PBS. Then, cells were incubated with annexin V-fluorescein in a HEPES buffer containing PI. The samples were analyzed in a FACSCalibur flow cytometer (BD Biosciences, Heidelberg, Germany). Ten thousand cells from each sample were analyzed, and both histogram and dual-parameter dot plot analyses were performed with the WinMDI 2.7 software. Detection of apoptotic cells by this method is based on Ca2+-dependent binding of annexin V-fluorescein to the negatively charged phospholipid surfaces of apoptotic cells and shows high specificity for phosphatidylserine. Instrument parameters used were: 488 nm for excitation, a 515-nm bandpass filter for fluorescein detection, and a filter >600 nm for PI detection.
For staining of apoptotic nuclei of HtTAM cells, cells were cultured in normal medium and TFM as for the FACS assay. Afterward, the cells were fixed in 5% formaldehyde, and apoptotic nuclei were stained with 0.05 mg/ml PI and preparations were examined with a fluorescence microscope. Apoptotic cells, as characterized by the morphology of their PI-stained nuclei (38) were counted, along with intact nonapoptotic nuclei. Percent apoptotic cells ± SEM was calculated as follows: number of apoptotic cells/total cells x 100 = percent apoptotic cells.
DNA fragmentation assay
In some of the experiments, the DNA fragmentation assay was performed to examine the degree of cellular apoptosis. Following treatment with cam, TFM, and rMIF or control buffer (see above), 1 x 106 cells were lysed in 1 ml of extraction buffer (SDS 1%, NaCl 0.4 M, EDTA 10 mM, Tris-hydrochloride, pH 8.0, 10 mM). After a 1-h incubation at 37°C, 10 µg/ml proteinase K was added and the samples were incubated for 3 h at 50°C. Samples were then extracted with phenol/chloroform/isoamyl alcohol (25:24:1) followed by a chloroform extraction, and precipitation of the DNA by a routine protocol. The DNA was resuspended in 40 µl of bidistilled water, 0.5 µl RNase A was added, and the solution was incubated for 45 min at 37°C. The DNA was then directly applied to a 1% agarose gel for electrophoretic separation and bands were visualized by ethidium bromide staining.
Measurement of GSH levels
Intracellular GSH levels were measured in lysates from cells overexpressing human MIF in a Tet-off-dependent manner.
For GSH determinations from HtTAM cell lysates, 2 x 106 cells were plated in 75-cm3 cell culture flasks and incubated for 72 h at 37°C in the presence vs absence of 1 µg/ml dox. Cells were washed once with PBS and twice with TFM. dox or control solution (1 µg/ml) was added and the cells were cultured in TFM for another 18 h at 37°C. Cells treated with 200 µM L-cystine served as a positive control. The cells were harvested and assayed for intracellular GSH by a commercial GSH assay (Bioxytech GSH-400 colorimetric assay; Novabiochem/Calbiochem). Briefly, the cells were removed from the flasks by incubation in calcium-magnesium-free-PBS and 4 x 106 cells were resuspended in 5% freshly prepared metaphosphoric acid (MPA). The suspensions were mixed by rigorous pipetting and subjected to dounce homogenization applying 20 pushes with a tight pestle. The resulting suspension was transferred to a 1.5-ml tube, centrifuged, and the supernatant was placed on ice. The GSH content of the acid-soluble supernatant was then measured in triplicate wells of a 96-well plate with 0.025% (w/v) lubrol according to the manufacturer’s instructions. The absorbance was recorded at 1-min intervals for 5 min at 405 nm.
For GSH determinations in Kym-1 cells, 2 x 106 cells were plated in 75-cm3 cell culture flasks, subjected to the transient transfection procedure with pMIFRP, and treated with dox as described above. Cells were washed with PBS and TFM and incubated in TFM as described for the HtTAM cells. The quantification of the cellular GSH content was performed as for the HtTAM cells, except that MPA was added directly to the cell layer. Obtaining homogeneous Kym-1 cell lysates was problematic for unknown reasons. Direct addition of MPA to Kym-1 cells followed by dounce homogenization was found to be the best method to get reliable GSH readings for these cells, while various other homogenization procedures including ultrasonification in combination with mechanical methods and ultraturrax treatment did not yield satisfactory results (data not shown).
MIF ELISA
The cellular MIF content following tet-induced overexpression was measured by a commercial ELISA from R&D Systems (Wiesbaden, Germany). The cell lysates to be subjected to the ELISA determinations were obtained applying the Tris-phosphate/CDTA/Triton lysis solution (see -Galactosidase assay) and lysates were diluted appropriately in the incubation buffer of the ELISA kit.
Analysis of the phosphorylation of endogenous c-Jun
For measuring the levels of endogenous phospho-c-Jun, 2 x 106 HtTAM cells were plated in 75-cm3 cell culture flasks and incubated for 72 h at 37°C in the presence vs absence of 1 µg/ml dox. Cells were then removed from the flasks and washed once with PBS and twice with TFM. Cells were resuspended in TFM and 1.5 x 106 cells were plated in a 3.5-cm dish and incubated for 2–6 h in the presence vs absence of 1 µg/ml dox. Cells were harvested, washed once with PBS, and lysed. Lysis was performed in RIPA buffer (1x PBS, pH 7.4, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS). The protein concentration in the cell lysates was determined by the Bradford method (39). Lysates were adjusted to contain equal protein concentrations and were mixed 1:1 with 2x Laemmli SDS-PAGE buffer. Electrophoresis was performed in 12% SDS-PAGE gels and separated proteins were transferred to a nitrocellulose membrane. Membranes were blocked with 1% BSA in TBST buffer (20 mM Tris-HCl, pH 7.6, 137 mM NaCl, 0.1% Tween 20 (v/v)) for 1 h at room temperature and then were incubated for 2 h at room temperature (or for 16 h at 4°C) with a monoclonal anti-phospho-c-Jun Ab (clone KM-1; Santa Cruz Biotechnology, Heidelberg, Germany) dissolved in TBST buffer plus 1% BSA followed by incubation with peroxidase-conjugated secondary Ab (Jackson ImmunoResearch Laboratories/Dianova, Hamburg, Germany) for 1 h at room temperature. Membranes were washed three times in TBST and immunoreactive bands were visualized using the Super Signal West Dura Extended Duration Substrate (Pierce/KMF Laborchemie, St. Augustin, Germany). For quantification, the blots were reprobed for c-Jun and actin staining. Membranes were stripped with 0.5% Tween 20/TBS for 1 h at room temperature, washed, and stained with an anti-c-Jun (clone H-79; Santa Cruz Biotechnology) or anti-actin (Sigma-Aldrich) Ab. The intensity of the phospho-c-Jun, c-Jun, and actin bands was quantified by densitometry using the Aida 2D software (Raytest Isotopenmessgeräte, Straubenhart, Germany).
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
One such system was obtained by stably transfecting the tet-regulated HeLa cell line HtTA with the cDNA for human MIF. HtTA cells contain significant baseline concentrations of MIF (123 ± 19 fg/cell; mean ± SD, n = 3), but preliminary experiments with transiently transfected tet-regulated MIF plasmids had suggested that good specific induction levels could be obtained with this system. Of a total number of 36 stable clones obtained, 3 were tested further and screened for their dox-dependent MIF expression levels. Clones 13 and 14 showed the most favorable combination of low baseline MIF expression values and high induction levels and had switch-on factors for the expression of MIF of 5.2 ± 0.2 and 2.6 ± 1.1, respectively (mean ± SD, n = 3) (Fig. 1A). Clone 13 exhibited a baseline MIF concentration of only 96 ± 26 fg/cell (mean ± SD, n = 3) in the presence of 1 µg/ml dox (switch-off status), which is <50% of the concentration of MIF measured in most other cell types (40). Clone 13 was thus used for the apoptosis experiments in the study. HeLa cells stably transfected with both the tet gene regulation cassette and the human MIF gene were designated HtTAM cells.
We used the antitumor drug and topoisomerase I inhibitor cam as a strong stimulus for the induction of apoptosis (41, 42). Overexpression of MIF in HtTAM cells initiated by the removal of dox from the culture medium protected cells from cam-induced apoptosis by up to 40%, depending on the concentration of cam applied (Fig. 1B). For comparison, when apoptosis was induced by growing the HtTAM cells to high density (cell density-induced apoptosis; Ref. 43), the degree of suppression of apoptosis yielded by the overexpression of MIF was 20% (n = 3; data not shown). We observed that the degree of protection provided by MIF following these stimuli, while significant, was only partial, ranging from 20 to 50% (see also below).
The rhabdomyosarcoma cell line Kym-1 was chosen as another experimental system. Kym-1 cells only contain minute concentrations of MIF protein, if any (none by Western blotting and 7 ± 5 fg/cell (mean ± SD, n = 3) by MIF ELISA; see Ref. 24), thus exhibiting a very favorable background expression level. We were not able to generate Kym-1 clones stably expressing both the tet regulator and human MIF genes (data not shown). However, transient overexpression of the tet and MIF plasmids in these cells led to excellent specific induction levels of MIF (Fig. 1C). When apoptosis in these cells was induced by growing the cells to high density, MIF led to a reduction of apoptosis by 30% (Fig. 1D).
Together, these data showed that intracellular MIF, following ectopic overexpression, was able to inhibit apoptotic processes induced by different stimuli. As these effects seemed to strictly depend on the enhancement of the concentration of intracellular MIF protein (Fig. 1, A and C), they may represent MIF activity that may not be related to its repertoire of activities of a transcellularly acting cytokine. Thus, we next tested whether biologically active rMIF added exogenously to cells could also inhibit apoptosis. The human promonocytic cell line HL-60 was applied for these experiments, as the monocyte/macrophage had been demonstrated to be the most important cellular target of the immunological and inflammatory cytokine activities of MIF (7, 44). cam led to a strong apoptotic response in these cells (Fig. 1E), confirming previous results by others (45). rMIF, when added to HL-60 cells for 24 h before cam treatment, markedly reduced the degree of apoptosis in a concentration-dependent manner. The protection from apoptosis provided by MIF ranged from a few percent (for 1 nM MIF) up to 50% (for 3 µM MIF) (Fig. 1E). The potential antiapoptotic activity of rMIF on immune cells was also probed in a cellular model of UV stress-induced apoptosis of T cells. We found that, when Jurkat T cells were exposed to UV irradiation for 1 or 6 h, preincubation with rMIF (2 µM) led to an inhibition of apoptosis of 20–30% (Fig. 1F).
Together, these experiments demonstrated that MIF could inhibit apoptosis induced by several stimuli in a variety of cell types. It appears that both intracellular MIF, possibly representing its role as an abundant enzyme and cell regulator, and exogenously added rMIF, representing the activities of MIF as a typical cytokine, can protect cells from apoptosis.
MIF has been shown to share several functional homologies with proteins of the Trx family of oxidoreductases (15, 46). Like Trx, MIF exhibits an enzymatic thiol-protein oxidoreductase activity that is dependent on the presence of a Cys-Xaa-Xaa-Cys (CXXC) redox motif (15, 21). Ample evidence is now available to show that Trx exhibits cytokine-like properties (47, 48, 49) and participates in the regulation of cellular redox homeostasis (18, 46, 49, 50, 51). Trx acts as a potent inhibitor of redox stress-induced apoptosis and this activity was found to be dependent on the presence of cysteine residues of Trx (51, 52, 53). These similarities between MIF and Trx prompted us to investigate whether MIF may also play a role in redox stress-induced apoptotic events. As the oxidoreductase enzymatic activity of MIF had been demonstrated to depend on the presence of the CALC sequence motif of MIF and, in particular, on the presence of cysteine residue 60 (Cys60), we first compared the apoptosis-regulating properties of wtMIF with those of the enzyme-dead mutant C60SMIF. Induction of C60SMIF was comparable to that of wtMIF (Fig. 1C). When the apoptosis-regulating effect of overexpressed wtMIF in Kym-1 cells was compared with that of the cysteine mutant, we observed that overexpression of C60SMIF did not lead to any suppression of apoptosis (Fig. 2A).
|
To further establish a potential role for MIF as an inhibitor of redox stress-induced apoptosis as suggested by our findings in Fig. 2A, we directly studied the role of MIF in pro-oxidative stress-induced apoptosis in HL-60, HtTAM, and HtTA1–1 cells.
Thiol starvation as a stimulatory event for mild oxidative stress was applied first to induce apoptosis to address this question. Annexin V/PI-based FACS analysis of HL-60 cells showed that a significant portion of up to 40% of the cells undergoes apoptosis following TFM treatment, whereas only little necrosis was observed under these conditions (Fig. 3A). A potential protective effect of MIF was then compared with the well-known protective effects of small molecule thiol compounds such as 2-ME and L-cys. We found that rMIF added at a concentration of 2 µM led to an inhibition of thiol starvation-induced apoptosis of 30%. For comparison, 2 µM 2-ME reduced apoptosis by 50% and 200 nM L-cys rescued 15% of the starved HL-60 cells (Fig. 3B). These data demonstrated that MIF was able to protect cells from pro-oxidative stress-induced apoptosis. As concentrations of MIF below 1 µM did not exhibit a significant antiapoptotic effect (data not shown), these data also suggested that a substantial amount of MIF-derived thiol equivalents was necessary for this effect to occur. A similar observation has been made for Trx (52).
|
|
20% in thiol-starved Jurkat T cells, when MIF was transiently expressed in these cells by tet gene regulation (data not shown). When apoptosis was induced by cell density stress or cam, no significant effect of MIF on the cytosolic levels of GSH could be observed, neither did rMIF further enhance the levels of cellular GSH in HtTA cells, once intracellular MIF was overexpressed (data not shown). Diamide treatment has been shown to also represent a valuable model of redox stress induction (56). Therefore, we tested next whether apoptosis in HtTAM cells was inducible by low concentrations of diamide and could be reversed by induction of MIF. Treatment of HtTAM cells with 0.2 mM diamide for 20 min induced marked apoptosis in these cells and apoptosis was significantly inhibited by ectopic overexpression of MIF (Fig. 5A). A time course analysis at this diamide concentration indicated that diamide-induced apoptosis increased up to at least 2 h of treatment. By contrast, apoptosis in the MIF-overexpressing cells did not increase further beyond 50 min, indicating that the maximal protective effect of MIF occurs in the range of 1–2 h following stimulation. With respect to these parameters, the diamide model differs from the thiol starvation model, as apoptosis induction and reversal by MIF in that system was not observed until 6 h after treatment (data not shown).
|
|
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Over the past years, MIF has been recognized as a pluripotent immunomodulator with broad inflammation-regulating properties including stimulatory effects on cell proliferation and cellular activation (i.e., summarized in Refs. 3 and4). Moreover, it appears that MIF could be one of the few target proteins identified which could serve as a connection between innate immunity and cancer (29, 34). However, the precise molecular changes induced by MIF in such situations have not yet been fully elucidated. Initial studies have indicated that activation of cell proliferation by MIF could be due to a bypassing of p53-mediated growth arrest. It has been proposed that this functional inactivation of p53 by MIF is based on the suppression of p53-dependent transcriptional activity by MIF (29). Mitchell et al. (34) found that MIF-mediated inhibition of p53 requires cyclooxygenase activity. Stimulation of the extracellular signal-regulated kinase 1/2 mitogen-activated protein kinase activity was identified as a potential pathway by which MIF modulates cell proliferation (26). In contrast, other studies have indicated that MIF can potently inhibit cell proliferation in fibroblasts (24), that it may decrease CD8+ T cell life span (28), and that it may inhibit the proliferation of microvascular endothelial cells (R. Kleemann and J. Bernhagen, manuscript in preparation). Furthermore, MIF acts to antagonize JAB1/COP9 signalosome (CSN)5-mediated rescue of fibroblasts from starvation-induced growth arrest (24).
It has been suggested that MIF could be a critical mediator of tumorigenesis (3, 57). This notion has mainly been based on the observed correlation of enhanced MIF expression levels and tumor occurrence as well as on the potent antitumor activity of anti-MIF Ab treatment (3, 57). As the anti-tumor activity of anti-MIF Abs appears to be due to an inhibition of angiogenic processes (57) and because in some models, MIF expression inversely correlates with tumor progression (11), the role of MIF in tumorigenesis is probably not simply due to an enhancement of cell proliferation. Rather, the effects of MIF on cell proliferation, growth arrest, cell cycle, and apoptotic processes need to be carefully discerned.
As to a contribution of MIF to the regulation of cell survival, the results of the present study together with the previous data by Hudson et al. (29) and Mitchell et al. (34) unanimously suggest that MIF acts to suppress apoptosis. Upon first sight, the inhibitory effect of MIF on apoptosis correlates with its reported stimulatory effects on cell proliferation. However, cellular apoptosis and cell proliferation are processes that are regulated in a very complex fashion and that do not simply oppose each other. Cytokines are typical participants in this regulatory network. In fact, several cytokines have been shown to be able to both promote and inhibit apoptosis and cell proliferation, depending on the cell type, the specific cellular activation state, and the physiological context (58, 59). It is thus not surprising that the observed apoptosis-suppressing effects of MIF are not complete. We found that inhibition of apoptosis by MIF induced by most stimuli applied ranged from 20 to 60% with a higher extent of inhibition seen, when apoptosis was induced by redox stress. Hudson et al. (29) have obtained similar results; they have observed that in starvation and estradiol treatment-induced apoptosis of fibroblasts, MIF reduced cell death by a margin of 20%, while NO radical-induced apoptosis of macrophages is much more drastically suppressed by MIF (up to 70%) (29, 34). Together with our results, the latter data suggest that suppression of apoptosis by MIF is most profound when apoptosis is caused by oxidative (or nitrosative) stress.
The role of MIF in apoptosis is closely associated with its inhibitory effects on p53 function, with three downstream targets of p53-dependent transcriptional activation, namely p21Cip1, cyclin G, and MDM2, identified to be regulated by MIF and p53 (29). In part, this activity could be due to a MIF-mediated inhibition of p53 and phospho-p53 accumulation (34). Our finding that MIF suppresses pro-oxidative stress-induced apoptosis, in addition to representing a potential independent pathway, indicates that MIF could alter p53 activity by a redox mechanism. Such a scenario appears plausible because redox regulation has been shown to be one of the pathways critical for cell viability (49, 50, 51). Moreover, the cocytokine Trx has been demonstrated to directly suppress apoptosis by virtue of its redox-regulating activity (49, 52, 53). p53 function is based on the zinc-mediated sequence-specific DNA binding activity of p53, which is dependent on at least three reduced cysteine residues. Oxidation of these cysteines can lead to conformational changes in the p53 DNA binding domain, resulting in impaired DNA binding activity. Thus, p53 activity is potently and directly regulated by redox-based mechanisms (60, 61). Trx, but also redox factor-1, another cellular redox stress protein (62) that modulates transcription factor activity by keeping critical cysteine residues of these proteins in a reduced state, have been implicated to participate in sustaining p53 conformation and activity through their reducing properties (61). By analogy, MIF might therefore also be capable of reducing the cysteine thiol groups of p53, conferring protection from cysteine oxidation during redox stress. Two additional lines of evidence support the notion that p53 activity could be regulated by MIF-mediated redox processes. First, MIF was recently shown to interact with the transcriptional coactivator JAB1/CSN5 and to modulate the cell cycle through JAB1 (24). At least some of the JAB1-modulating activities of MIF appear to be connected to the presence of the redox-active cysteine 60 residue of MIF. Secondly, JAB1 and p53 have been demonstrated to specifically interact with each other (35). Together, these observations offer the possibility that a ternary MIF-JAB1-p53 complex could be the molecular basis for MIF-mediated, p53-dependent suppression of apoptosis. However, neither has complex formation between MIF and p53 been demonstrated, nor has an involvement of MIF in p53-JAB1 complexes yet been investigated.
It is also speculative at this time whether redox regulation of apoptosis by MIF occurs by redox catalysis or whether the redox-competent cysteine thiol groups of MIF serve to enhance the cell’s pool of reducing thiol equivalents. The latter possibility is supported by initial data from experiments with MIF-overexpressing HtTAM cells that were additionally treated with rC60SMIF and that indicated that rC60SMIF led to a small decrease in apoptosis compared with rMIF-treated control cells. This could mean that residues Cys57 or Cys81 may serve as an additional thiol pool under these conditions. Interestingly, Yodoi and coworkers (52) performed a detailed investigation of the role of the redox-active cysteine residues of Trx in the apoptosis of T cells. Applying various cysteine mutants of Trx, they found that Trx-mediated suppression of redox stress-induced apoptosis is dependent on the presence of the cysteine thiols. As also mutant C31S/C34STrx, with both CXXC cysteines changed, as well as partially oxidized Trx, suppressed pro-oxidative stress-induced apoptosis to some extent, it appears that Trx-type oxidoreductases modulate cell viability by both CXXC-dependent catalysis, but also by direct thiol group donation that may also be mediated by the non-CXXC cysteine residues. In fact, increasing evidence suggests that cellular redox homeostasis is regulated not only by the ratios of oxidized vs reduced small molecule thiols such as GSH, but that oxidoreductases such as Trx directly participate in regulating the cellular redox state by "donation" of proteinaceous thiol equivalents (51, 52).
A membrane receptor for MIF has not yet been identified, but evidence suggests that a receptor may exist (26, 54). Both receptor-mediated and intra/autocrine pathways of MIF action have been discussed and may occur in parallel (54). Our finding that the redox-dead mutant C60SMIF strongly suppresses apoptosis, when applied exogenously, but is inactive, when overexpressed endogenously, could imply that the redox activity of MIF acts differently on the potential target proteins. Such target activities might include binding to PAG or redox regulation of JAB1 and p53. However, as different apoptotic stimuli were used in the experiments with endogenous overexpression as compared with exogenous addition of recombinant protein (i.e., density stress and cam treatment, respectively), this conclusion certainly needs to be broadly based on further studies. Nevertheless, an intracellular role for the redox activity of MIF is also suggested by the finding that the redox stress-treated HtTAM cells showed a lower degree of JNK activity when MIF was induced. Of note, JAB1-containing CSN function was recently linked to JNK and p53 activity (reviewed in Ref. 63). The observed kinetics of the induction of phospho-c-Jun by TFM treatment and inhibition by MIF are consistent with the suggested role for JNK as an upstream stimulatory event during stress-induced apoptosis (64).
The similarities between MIF and Trx in the regulation of apoptosis add to the previously noted functional homologies between MIF and Trx-type proteins (15). For example, the redox activity of MIF has been implicated to be important for the interaction between MIF and PAG (22). Likewise, interaction of Trx and redox factor-1 has been found to be redox-dependent (65). Also, the various immunoregulatory activities of MIF and Trx have been shown to be at least partially dependent on the presence of the corresponding redox-active cysteine residues (15, 47). It is noteworthy to mention that Trx has recently been redefined as a cocytokine (47, 48, 49).
For MIF, a correlation between its redox activities and its functions in immune regulation is also becoming increasingly evident. For example, MIF redox activity is at least in part critical for the glucocorticoid-overriding activity of MIF (21). Furthermore, MIF functions as a redox-sensitive cytokine in myocardial ischemia (23).
In summary, modulation of redox stress-induced apoptosis by MIF could be an important novel mechanism, linking the redox function of MIF and its role in immune and stress regulation.
|
Acknowledgments |
|---|
|
Footnotes |
|---|
2 Address correspondence and reprint requests to Dr. Jürgen Bernhagen, Department of Biochemistry and Molecular Cell Biology, Institute of Biochemistry, University Hospital Rheinisch-Westfälische Technische Hochschule, Pauwelsstrasse 30, D-52074 Aachen, Germany. E-mail address: jbernhagen{at}ukaachen.de
3 Abbreviations used in this paper: MIF, macrophage migration inhibitory factor; CALC, Cys57-Ala58-Leu59-Cys60; Trx, thioredoxin; PAG, peroxiredoxin; JAB1, Jun activation domain binding protein 1; JNK, c-Jun N-terminal kinase; tet, tetracycline; wt, wild type; GSH, glutathione; L-cys, L-cysteine; cam, camptothecin; dox, doxycycline; TFM, thiol-free medium; CSN, COP9 signalosome; HtTA, tet-regulated HeLa cell; FM, full medium; PI, propidium iodide; MPA, metaphosphoric acid; CXXC, Cys-Xaa-Xaa-Cys.
Received for publication February 11, 2002. Accepted for publication January 6, 2003.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
cell macrophage migration inhibitory factor. Proc. Natl. Acad. Sci. USA 94:4782.1 and overexpression of Bcl-2. J. Biol. Chem. 272:20003.
on apoptosis of human neutrophils. J. Leukocyte Biol. 69:467.This article has been cited by other articles:
|
|
 |
|
  N. L. Millar, A. Q. Wei, T. J. Molloy, F. Bonar, and G. A. C. Murrell Cytokines and apoptosis in supraspinatus tendinopathy J Bone Joint Surg Br, March 1, 2009; 91-B(3): 417 - 424. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. L. Damico, A. Chesley, L. Johnston, E. P. Bind, E. Amaro, J. Nijmeh, B. Karakas, L. Welsh, D. B. Pearse, J. G. N. Garcia, et al. Macrophage Migration Inhibitory Factor Governs Endothelial Cell Sensitivity to LPS-Induced Apoptosis Am. J. Respir. Cell Mol. Biol., July 1, 2008; 39(1): 77 - 85. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M Mihm, P J Baker, L M Fleming, A M Monteiro, and P J O'Shaughnessy Differentiation of the bovine dominant follicle from the cohort upregulates mRNA expression for new tissue development genes Reproduction, February 1, 2008; 135(2): 253 - 265. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. Sun, H. Li, Y. Gao, T. Matsuura, P. A. Upchurch, M. K. Raizada, and C. Sumners Lack of Macrophage Migration Inhibitory Factor Regulation Is Linked to the Increased Chronotropic Action of Angiotensin II in SHR Neurons Hypertension, March 1, 2007; 49(3): 528 - 534. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. Schmidt, J. Hoffend, A. Altmann, F. Kiessling, L. Strauss, D. Koczan, W. Mier, M. Eisenhut, R. Kinscherf, and U. Haberkorn Troponin I Overexpression Inhibits Tumor Growth, Perfusion, and Vascularization of Morris Hepatoma J. Nucl. Med., September 1, 2006; 47(9): 1506 - 1514. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
P. Kunz, J. Hoffend, A. Altmann, A. Dimitrakopoulou-Strauss, D. Koczan, M. Eisenhut, G. A. Bonaterra, T. J. Dengler, W. Mier, U. Haberkorn, et al. Angiopoietin-2 Overexpression in Morris Hepatoma Results in Increased Tumor Perfusion and Induction of Critical Angiogenesis-Promoting Genes J. Nucl. Med., September 1, 2006; 47(9): 1515 - 1524. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 E. J. Beswick, I. V. Pinchuk, G. Suarez, J. C. Sierra, and V. E. Reyes Helicobacter pylori CagA-Dependent Macrophage Migration Inhibitory Factor Produced by Gastric Epithelial Cells Binds to CD74 and Stimulates Procarcinogenic Events. J. Immunol., June 1, 2006; 176(11): 6794 - 6801. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. Matsuura, C. Sun, L. Leng, A. Kapurniotu, J. Bernhagen, R. Bucala, A. E. Martynyuk, and C. Sumners Macrophage Migration Inhibitory Factor Increases Neuronal Delayed Rectifier K+ Current J Neurophysiol, February 1, 2006; 95(2): 1042 - 1048. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. Yao, S. Shida, M. Selvakumaran, R. Zimmerman, E. Simon, J. Schick, N. B. Haas, M. Balke, H. Ross, S. W. Johnson, et al. Macrophage Migration Inhibitory Factor Is a Determinant of Hypoxia-Induced Apoptosis in Colon Cancer Cell Lines Clin. Cancer Res., October 15, 2005; 11(20): 7264 - 7272. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Schmidt, J. Hoffend, A. Altmann, L. G. Strauss, A. Dimitrakopoulou-Strauss, B. Engelhardt, D. Koczan, J. Peter, S. Vorwald, H. Eskerski, et al. Transfer of the sFLT-1 Gene in Morris Hepatoma Results in Decreased Growth and Perfusion and Induction of Genes Associated with Stress Response Clin. Cancer Res., March 15, 2005; 11(6): 2132 - 2140. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. Sun, H. Li, L. Leng, M. K. Raizada, R. Bucala, and C. Sumners Macrophage Migration Inhibitory Factor: An Intracellular Inhibitor of Angiotensin II-Induced Increases in Neuronal Activity J. Neurosci., November 3, 2004; 24(44): 9944 - 9952. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. Eickhoff, C. Baldauf, H.-W. Koyro, G. Wennemuth, Y. Suga, J. Seitz, R. Henkel, and A. Meinhardt Influence of macrophage migration inhibitory factor (MIF) on the zinc content and redox state of protein-bound sulphydryl groups in rat sperm: indications for a new role of MIF in sperm maturation Mol. Hum. Reprod., August 1, 2004; 10(8): 605 - 611. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Suzuki, Y. Takamura, M. Maeno, S. Tochinai, D. Iyaguchi, I. Tanaka, J. Nishihira, and T. Ishibashi Xenopus laevis Macrophage Migration Inhibitory Factor Is Essential for Axis Formation and Neural Development J. Biol. Chem., May 14, 2004; 279(20): 21406 - 21414. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. T. Nguyen, J. Beck, H. Lue, H. Funfzig, R. Kleemann, P. Koolwijk, A. Kapurniotu, and J. Bernhagen A 16-Residue Peptide Fragment of Macrophage Migration Inhibitory Factor, MIF-(50-65), Exhibits Redox Activity and Has MIF-like Biological Functions J. Biol. Chem., September 5, 2003; 278(36): 33654 - 33671. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. A. Amin, O. V. Volpert, J. M. Woods, P. Kumar, L. A. Harlow, and A. E. Koch Migration Inhibitory Factor Mediates Angiogenesis via Mitogen-Activated Protein Kinase and Phosphatidylinositol Kinase Circ. Res., August 22, 2003; 93(4): 321 - 329. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
