Native, not nitrated, cytochrome c and mitochondria-derived hydrogen peroxide drive osteoclast apoptosis

Merry Jo Oursler,1,2 Elizabeth W. Bradley,2 Sarah L. Elfering,3 and Cecilia Giulivi2,3

Departments of 1Biology, Medical Microbiology and Immunology, 2Biochemistry and Molecular Biology, and 3Chemistry, University of Minnesota, Duluth, Minnesota

Submitted 17 February 2004 ; accepted in final form 26 August 2004


    ABSTRACT
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
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Two unresolved aspects of the role of mitochondria-derived cytochrome c in apoptosis are whether there is a separate pool of cytochrome c within mitochondria that participates in the activation of apoptosis and whether a chemically modified cytochrome c drives apoptosis. These questions were investigated using osteoclasts, because they are rich in mitochondria and because osteoclast apoptosis is critical in bone metabolism regulation. H2O2 production was increased during culture, preceding cytochrome c release; both processes occurred anterior to apoptosis. With the addition of a mitochondrial uncoupler, H2O2 production and apoptosis were blocked, indicating the prominent role of mitochondria-derived H2O2. Trapping H2O2-derived hydroxyl radical decreased apoptosis. Cytosolic cytochrome c was originated from a single mitochondrial compartment, supporting a common pool involved in respiration and apoptosis, and it was chemically identical to the native form, with no indication of oxidative or nitrative modifications. Protein levels of Bcl-2 and Bc-xL were decreased before apoptosis, whereas expression of wild-type Bcl-2 repressed apoptosis, confirming that cytochrome c release is critical in initiating apoptosis. Cytosolic cytochrome c participated in activating caspase-3 and -9, both required for apoptosis. Collectively, our data indicate that the mitochondria-dependent apoptotic pathway is one of the major routes operating in osteoclasts.

reactive oxygen species; nitric oxide; free radicals; caspase


OSTEOCLASTS ARE MULTINUCLEATED hematopoietically derived cells responsible for most, if not all, cell-based bone destruction in vertebrates. Osteoclast numbers are the main determinant of the rate of bone resorption, and it is of great interest to understand how osteoclast numbers are controlled (56). It is becoming increasingly evident that a decline in osteoclast number in vivo is the result of apoptosis (32, 44). Apoptosis is a controlled series of events that results in biochemical and morphological changes, including membrane blebbing, cell shrinkage, DNA fragmentation, chromatin condensation, and formation of apoptotic bodies. The mechanisms by which osteoclasts survive once they mature remain unresolved, but our recent studies have implicated phosphatidylinositol 3-kinase (PI3K) stimulation of both the AKT/NF-{kappa}B and the MEK/ERK pathways in osteoclast survival (14).

The release of cytochrome c from mitochondria to the cytosol is pivotal in the activation of caspases and the ensuing cell death (26). After a death stimulus, cytosolic Bax translocates to mitochondria (17, 67), where it can promote the release of cytochrome c (17, 23). Once released, cytosolic cytochrome c interacts with apoptotic protease-activating factor-1 (Apaf-1) in the presence of ATP, stimulating the processing of procaspase-9 to its active form, which in turn can then activate the executioner caspase-3 and caspase-7 (30, 38). Translocation of cytochrome c occurs in several experimental models of acute and chronic disorders. However, whether cytochrome c in osteoclasts has a role in the regulation of apoptosis is unresolved.

Oxygen- and nitrogen-reactive species have been associated with different stages of apoptosis. In this regard, it has been proposed that cytochrome c released from mitochondria could represent a hemoprotein chemically modified by oxidative/nitrative stress (19) or that released cytochrome c is originated from a microcompartment different from that of cytochrome c involved in the respiratory chain (2). We examined both of these possibilities in the studies reported in this article.

It was the goal of this study to understand the mechanisms of apoptosis in osteoclasts, focusing on the roles of mitochondria and cytochrome c in this process. We have shown that cytochrome c translocation occurred after an increase in hydrogen peroxide production by mitochondria was observed. We also have demonstrated that cytochrome c released from mitochondria is chemically identical to the native one and that it is not released from a different mitochondrial compartment (e.g., intermembrane vs. intracristae). These findings support the possibility that there is recruitment of the mitochondria-dependent apoptotic pathway in osteoclasts and indicate the critical role of hydrogen peroxide production by mitochondria as a key event in activating cell death.


    MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
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Chemicals and biochemicals. All reagents were of analytical grade. The spin trap 5,5'-dimethyl-1-pyrrolyne-N-oxide was purchased from Aldrich Chemical (Milwaukee, WI). The spin trap was purified by repeated charcoal filtration until its electron paramagnetic resonance (EPR) spectrum was virtually signal free. Unless otherwise noted, all chemicals were from Sigma (St. Louis, MO).

Osteoclast culture and purification. Mouse marrow cells containing osteoclast precursors were obtained from female BALB/c mice (Taconic, Germantown, NY) as previously described (14). Briefly, 4- to 6-wk-old mice were killed, long bones of the hindlimbs were aseptically removed, and the marrow cells were harvested. Marrow cells were cultured with ST2 stromal cells (Riken Cell Bank, Tsukuba, Japan) during differentiation as follows. For apoptosis assessment, ST2 cells in {alpha}-modified minimal essential medium ({alpha}MEM; GIBCO BRL, Grand Island, NY) supplemented with 10% fetal bovine serum (Hyclone, Logan, UT) and antibiotic/antimycotic were plated in 24-well plates on glass coverslips at a density of 1.6 x 105 cells/well. For biochemical assessments, ST2 cells were plated in six-well plates at a density of 6.4 x 105 cells/well. After 24 h of culture, marrow cell precursors were plated as follows: for 24-well plates, marrow cells were added at 1.18 x 105 cells/well, and for 6-well plates, marrow cells were added at 4.75 x 105 cells/well. Precursors were added to the stromal cells in base medium composed of MEM, 10% fetal bovine serum, 1% nonessential amino acids, and 1% penicillin/streptomycin. The base medium was supplemented with 7 x 10–3 M ascorbic acid (GIBCO BRL, Rockville, MD), 0.1 µM dexamethasone, and 10 µM vitamin D3 (Biomol, Plymouth Meeting, PA) immediately before use (57, 61). Cells were fed every 3 days until 13 days of culture. Osteoclast-like cells were purified by 15-min treatment at 37°C with 0.2 mg/ml collagenase (Worthington Biochemical, Lakewood, NJ) in Ham's F-12 medium (GIBCO BRL, Grand Island, NY) followed by 30-min treatment at 37°C with 0.2 mg/ml dispase (Boehringer Mannheim, Framingham, MA) in Ham's F-12 medium to remove support cells, as has been previously documented in our laboratory (14).

Apoptosis detection. Purified osteoclasts were cultured in base medium for the indicated time period, fixed with 1% paraformaldehyde, and stained. Staining for apoptotic cells was performed by using Hoechst stain to detect chromatin condensation, annexin V binding to detect externalization of phosphatidylserine, and the fragment end labeling kit FragEL from Calbiochem (La Jolla, CA) to detect DNA fragmentation (14). Briefly, fixed osteoclasts were stained for 60 min with Hoechst 33258 diluted to 5 µg/ml in phosphate-buffered saline (PBS) with 0.01% Tween 20. The cells were then tartrate-resistant acid phosphatase (TRAP) stained using a kit from Sigma. To examine the timing of osteoclast apoptosis, mature cells were purified as described in Osteoclast culture and purification, and culture continued for up to 48 h. Cells were fixed and then stained with Hoechst and TRAP as described above. The slides were numbered and prepared for a blinded apoptosis assessment. As previously described by our laboratory group (14), DNA fragmentation and chromatin condensation are virtually identical in their sensitivity for detecting apoptotic osteoclasts. We selected chromatin condensation for routine analysis because this technique is the more efficient method in our hands. Cells were examined with fluorescent microscopy, and cells displaying strongly labeled nuclei were scored as apoptotic.

Hydrogen peroxide production by osteoclasts. Hydrogen peroxide released from cultured, purified osteoclasts or freshly isolated hepatocytes (59) into the overlying medium (without phenol red) was assayed at various time points. Aliquots from the supernatants were used to evaluate the content of hydrogen peroxide by using the oxidation of p-hydroxyphenylacetic acid catalyzed by horseradish peroxidase (HRP) (48); the concentrations were plotted against time, and the slopes were taken as representative of the rates of production of hydrogen peroxide. The concentration of hydrogen peroxide used to plot standard curves was determined spectrophotometrically at 240 nm (43.6 M–1·cm–1).

Cytochrome c detection by Western blot analysis. Osteoclasts were purified and cultured for the indicated time period, followed by incubation at 4°C for 15 min without or with 1 µg of digitonin in 200 µl of PBS. Commercial digitonin was recrystallized according to the method described by Kun et al. (27). After incubation, the supernatant was carefully removed and concentrated using Centricom tubes with a molecular weight cutoff of 3,000. The concentrate (~20 µl) was mixed with sample buffer (50 mM Tris, 2% SDS, 0.1% bromphenol blue, and 10% glycerol, pH 6.8). Sample buffer (100 µl) was added to the attached cells, and the cells were scraped from the plate. The concentrate and cells were analyzed (20 µl of cells and total amount of concentrate) on 15% SDS-PAGE gels, and the gels were electroblotted to nitrocellulose membranes. The membranes were blocked for 1 h at room temperature in a solution constituted by 1% nonfat dry milk, 10% normal goat serum, and 2% Tween 20 in Tris-buffered saline (TBS-T). A monoclonal antibody for cytochrome c (kind gift of Prof. Ronald Jemmerson, University of Minnesota) was used to probe for the hemoprotein by diluting it (1:750) in blocking solution and incubating it with the membranes for 1 h at room temperature. After washing, the membranes were incubated with HRP-linked goat anti-mouse antibodies (1:10,000 in blocking solution). The blots were developed using chemiluminescence visualized by autoradiography. The films were analyzed using the NIH Image software, and the results were expressed as the ratio of cytochrome c in the supernatant (cytochrome c released in the supernatant by digitonin treatment) to the total (no digitonin). Similar results were obtained by expressing the results as released cytochrome c per cytochrome c pellet.

Nitric oxide synthase detection by Western blot analysis. Six-well plates of purified osteoclast cultures, differentiated and purified as described in Osteoclast culture and purification, were washed with PBS, and then each well was lysed in 100 µl of 2x Laemmli buffer with 2-mercaptoethanol. The samples were heated for 5 min at 95°C, and 20 µl were loaded per lane of 7.5% SDS-PAGE gels. Gels were transferred to polyvinylidene difluoride (PVDF) membranes for 1 h and 15 min in transfer buffer containing 5% methanol, 48 mM Tris base, 39 mM glycine, and 1.3 mM SDS. Membranes for endothelial (eNOS) and inducible nitric oxide synthase (iNOS) were blocked in 10% normal goat serum, 1% nonfat dry milk, and 1% BSA in TBS-T for 1 h at room temperature. Membranes for neuronal NOS (nNOS) were blocked in 5% nonfat dry milk in TBS-T. Mouse monoclonal antibodies for eNOS and iNOS (BD Transduction Laboratories, San Diego, CA) were diluted at 1:2,000 in blocking buffer, and rabbit polyclonal antibodies for nNOS (Santa Cruz Biotechnology, Santa Cruz, CA) were diluted at 1:500 in blocking buffer. All primary antibody incubations occurred overnight. Secondary antibodies conjugated to HRP, and chemiluminescence reagents were used to detect protein bands. All blots were visualized on a Kodak Image Station 1000. PVDF membranes from the eNOS, iNOS, and nNOS Western blots were stripped, blocked in 5% nonfat dry milk, and reprobed with anti-actin goat polyclonal antibody (Santa Cruz Biotechnology) at 1:100 dilution overnight at 4°C. Bands were detected with rabbit anti-goat-HRP (Santa Cruz Biotechnology) at 1:5,000 dilution and visualized on a Kodak Image Station 1000.

Matrix-assisted laser desorption ionization with time-of-flight spectrometry of cytochrome c. Pellets and supernatants from osteoclasts cultured and purified as described in Osteoclast culture and purification were pooled and separated on 4–20% SDS-PAGE gels under reducing conditions. Protein bands were visualized with Coomassie blue, and parallel Western blots were used to identify the band corresponding to cytochrome c. This band was excised, crushed, and destained by extensive washing with 50% acetonitrile and 25 mM NH4CO3, pH 8.0. Protein was reduced in the gel with 50 µl of 50 mM diethyldithiothreitol (DTT) in 100 mM NH4CO3, pH 8.0, for 30 min at 56°C, and then the DTT solution was removed and replaced with 50 µl of 100 mM iodoacetamide in 100 mM NH4CO3, pH 8.0, for 30 min at 45°C. The alkylating solution was removed, and the gel pieces were washed once with 100 mM NH4CO3, pH 8.0, and then with acetonitrile and then dried in a Speed Vac centrifuge. Dried gel pieces were swollen in a minimal amount of trypsin (33 µg/ml; Promega) in 50 mM NH4CO3, pH 8.0, and digested overnight at 37°C. After digestion, peptides were extracted with 100 mM Na2CO3, pH 10, for 1 h at 37°C. Peptide extracts were desalted on Zip tips (Millipore, Bedford, MA) by being washed with 0.1% formic acid and eluted with 50% acetonitrile and 0.1% formic acid, as suggested by the manufacturer.

Peptides generated from the tryptic digestion of pellet or supernatant cytochrome c were separated by liquid chromatography connected to an electrospray mass spectrometer. Full scans of the samples from 600 to 3,500 m/z and tandem mass spectral data of select ions were collected on a QSTAR Pulsar i (Applied Biosystems, Foster City, CA) quadrupole time-of-flight mass spectrometer with an orthogonal matrix-assisted laser desorption ionization (MALDI) source by using dihydroxybenzoic acid as matrix (Agilent Technologies, Palo Alto, CA). External calibration was performed using human angiotensin II (monoisotopic [MH+] m/z 1046.5; Sigma) and adrenocorticotropic hormone (ACTH) fragment 18–39 (monoisotopic [MH+] m/z 2465.2; Sigma). All mass spectrum analyses were performed at the Mass Spectrometry Facility at the University of Minnesota, St. Paul campus. Peptide masses were searched by using the mass spectrum fit component of the Protein Prospector platform as well as ProFound software. For all digests, the oxidized Met modification was selected as well as the alkylation of Cys residues.

Gene array analysis of Bcl-2 family genes. Mouse Q Series apoptosis gene arrays (SuperArray, Bethesda, MD) were used to compare transcript levels of several Bcl-2 family genes. Total RNA samples were isolated from purified osteoclasts by using Trizol reagent (Invitrogen, Carlsbad, CA). A 1-mg sample of total RNA was reverse transcribed with MMLV reverse transcriptase (Roche, Indianapolis, IN) in the presence of biotin-16-dUTP (Roche). The resulting cDNA was hybridized to each array, and chemiluminescence detection was performed according the manufacturer's specifications. The signal from each Bcl-2 family gene hybridization was captured via autoradiography. Signals were converted into raw data with the use of ScanAlyze software (Michael Eisen, Lawrence Berkley National Lab, www.microarrays.org/software.html). Each raw data signal was then translated into expression levels by using GEArray Analyzer software. Expression levels of each gene were normalized to glyceraldehyde-3-phosphate dehydrogenase, and background subtraction was accomplished. Expression levels were determined in two replicate experiments to ensure accuracy.

Bcl-2 family member Western blot analysis. Osteoclasts were purified and either harvested immediately for Western blotting or cultured in base medium for the indicated time period, rinsed with PBS, and then harvested for Western blotting. Harvesting was accomplished by scraping into Laemmli sample buffer lacking {beta}-mercaptoethanol and bromphenol blue. To ensure that equal cell protein was analyzed, protein was determined using Bio-Rad's Protein Quantitation in Detergent Analysis kit. After protein quantitation, {beta}-mercaptoethanol and bromphenol blue were added to the samples, and 40 µg of protein were loaded in each lane. Parallel Western blotting was carried out as directed in the product literature by using antibodies directed against Bcl-2 (1:300 dilution; Oncogene Research Products), Bcl-xL/S (1:1,000 dilution; Cell Signaling), or tubulin (clone E7, 1:500 dilution; Developmental Studies Hybridoma Bank) and secondary antibodies (1:10,000) with chemiluminescence detection (Pierce, Rockford, IL). Antibodies were from Oncogene Sciences (San Diego, CA).

Infection of osteoclasts with Bcl-2 adenoviral expression vector. The Bcl-2 adenoviral expression vector was a gift from Dr. Hanjoong Jo (University of Alabama, Birmingham, AL). Recombinant viruses were produced in HEK-293 cells, purified, and titrated according to standard methods (22). Osteoclasts were differentiated in either 24-well plates with coverslips for apoptosis analysis or 6-well plates for protein expression analysis. Purified osteoclasts were infected with recombinant virus or empty vector-containing virus at multiplicities of infection (MOI) of 1 and 8 per nucleus in culture medium. After 24 or 48 h, cells were washed once with PBS. Cells on coverslips were fixed, stained, and analyzed as described in Bcl-2 family member Western blot analysis, and cells in 6-well plates were processed for Bcl-2 and tubulin by performing Western blotting as described.

EPR spectroscopy. EPR spectra were recorded at 9.2 GHz on a Varian E-102 spectrometer. Measurements were carried out with 100-kHz field modulation at room temperature. Purified osteoclasts were supplemented at time 0 with 200 mM 5,5'-dimethyl-1-pyrrolyne-N-oxide (DMPO), and aliquots of the supernatants were withdrawn at various time points. These aliquots were transferred to flame-sealed capillary ends of Pasteur pipettes. Instrument settings were as follows: receiver gain, 1 x 104; microwave power, 20 mW; microwave frequency, 9.22 GHz; modulation amplitude, 0.5 G; center field, 3,481 G; sweep width, 100 G; time constant, 1 s; scan time, 8 min/each spectrum for a maximum of five scans.

Statistical analysis. Results represent the means ± SE of three replicates from one experiment. Each experiment was carried out a minimum of three times, and the results shown are representative of all results obtained. The effect of treatment was compared with control values by using one-way analysis of variance (ANOVA); significant treatment effects were further evaluated by using the Fisher's least-significant difference method of multiple comparisons in a one-way ANOVA. Tests were carried out using StatView II (Abacus Concepts, Cupertino, CA) and StatSimple version 2.0.5 (Nidus Technologies, Toronto, Canada).


    RESULTS AND DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
 GRANTS
 REFERENCES
 
Kinetics of osteoclast apoptosis. Osteoclast survival was examined in culture after support cells were removed at day 13. The method used to separate support cells from osteoclasts resulted in multinucleated cells with TRAP-positive staining (Fig. 1). These two features (multinucleated cells and TRAP staining) are characteristic of mature osteoclasts, indicating that the purification process had effectively enriched the preparation with these cells. After 90 min of purification, mature osteoclast cultures contained both apoptotic (Fig. 1, circles) and surviving cells (Fig. 1, arrows) as evaluated by chromatin condensation, annexin V binding, and detection of DNA fragmentation.



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Fig. 1. Evaluation of osteoclast apoptosis. Purified osteoclasts were stained with tartrate-resistant acid phosphatase (TRAP) or stained for apoptosis as follows: chromatin condensation was evaluated using Hoechst stain, externalization of membrane phosphatidylserine was determined using annexin V binding, or DNA fragmentation was detected using the FragEl kit, as indicated. A: viable cells contained larger nuclei (circles), whereas apoptotic cells were more condensed (arrows). In some cells, as shown in B, there were nuclei within an apoptotic cell that stained for chromatin condensation yet were negative on the basis of DNA fragmentation. Images were acquired using a Nikon Eclipse E400 microscope equipped with a Sony charge-coupled device color video camera (model DCX-970MD). The images were processed using Adobe Photoshop.

 
Studies by our group (14) have shown that nuclear condensation, annexin V binding, and DNA fragmentation analyses are highly correlative, and we therefore selected chromatin condensation for our routine analyses. In some cells, as indicated in Fig. 1B, there are nuclei within an apoptotic cell that stain for chromatin condensation yet are negative on the basis of DNA fragmentation (arrows). However, we did not observe cells that were scored as apoptotic according to one detection method that were not likewise scored as apoptotic according to another detection method. In our studies, the nuclei in each multinucleated cell were uniformly either all uncondensed or condensed; thus there was no evidence of an intermediate phenotype. In some but not all experiments, we observed some apparent evidence that a few osteoclasts had undergone necrosis, but the frequency of this occurrence was independent of any treatment or further culture beyond the development of maturity. We attempted flow cytometry for further analysis of osteoclasts and found that the relatively large size (100 µm or larger) and difficulty of removing the cells from the culture plastic precluded any accurate assessment with this approach. The increase in apoptotic cells was significant after 90 min of culture (Fig. 2), confirming earlier studies of this timing (14).



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Fig. 2. Time course of osteoclast apoptosis and release of cytochrome c. Apoptotic osteoclasts (control, {circ}) were counted at the indicated intervals. Other experimental details were as described in Fig. 1. Cytochrome c release ({bullet}), expressed as the ratio of hemoprotein in the supernatant to the total content ([Cyt c]supernatant/[Cyt c]total), was evaluated at indicated time points according to the procedure described in MATERIALS AND METHODS. *P < 0.05 compared with time 0.

 
Role of mitochondria in osteoclast apoptosis. Efflux of cytochrome c from mitochondria is a pivotal event in apoptosis as it drives the assembly of a high-molecular-weight caspase-activating complex in the cytoplasm, termed the mitochondrial apoptosome (6, 43, 46, 70). The binding of cytochrome c to Apaf-1 is required for the activation of caspase-9 (32) in the presence of other essential factors such as ATP. To investigate the role of mitochondria in osteoclast apoptosis, we treated osteoclasts with digitonin under conditions that selectively lysed plasma membranes, whereas the mitochondrial ones remained intact. This method is based on the difference in chemical composition (i.e., cholesterol content; Ref. 66) of the plasma and mitochondrial membranes of cells. Experiments performed with osteoclasts aimed at optimizing the concentration of digitonin indicated that 0.01 mg digitonin/mg protein was required to lyse the plasma membrane, whereas a concentration five times higher was needed to lyse the outer mitochondrial membrane. These digitonin-to-protein ratios were within the range expected for other mitochondria-rich cells, e.g., hepatocytes (13, 71), and were identical for control and apoptotic cells. This latter observation precluded the existence of different pools of cytochrome c localized at various mitochondrial compartments (e.g., intracristae vs. intermembrane space) during the progression of apoptosis, which would have resulted in the need for different amounts of digitonin to release the cytochrome c.

Cytosolic (osteoclast supernatants), mitochondria-bound (pellets of osteoclasts), and total cytochrome c content (obtained as the addition of cytosolic and pellet-bound or as the total amount of cytochrome c released upon complete cell lysis) was evaluated by performing Western blots, using monoclonal antibodies to the hemoprotein. The amount of cytochrome c was evaluated by densitometry of the corresponding Western blots obtained at timed intervals. The release of cytochrome c increased linearly with time, in parallel to but preceding apoptosis detection (Fig. 2). Cytochrome c is present at relatively high concentrations in mitochondria (from 2.5 to 5 mM, considering an average concentration of cytochrome c of 0.2–0.5 nmol/mg mitochondrial protein and a volume of 81 µl/g mitochondrial protein; Ref. 54) and its release to the cytosol constitutes a critical event for the activation of caspases.

Osteoclasts are rich in mitochondria, and given that these organelles are the main source of reactive oxygen species (ROS) in vivo (15), it could be predicted that these cells may have a higher rate of hydrogen peroxide production. This assumption was confirmed by measurements of the rate of hydrogen peroxide production by osteoclasts (Table 1). Rates of hydrogen peroxide production by purified osteoclast cultures with and without FCCP indicated that 80–90% of the rate was originated by mitochondria (Table 1), for it is well known that in the presence of FCCP, an uncoupler, the mitochondrial contribution is negligible (15). The rate of hydrogen peroxide production was even higher than that of other mitochondria-rich cells, e.g., hepatocytes (Table 1). Evaluation of the steady-state concentration of hydrogen peroxide also indicated an almost threefold higher level than that found in hepatocytes (Table 1). It should be noted that primary hepatocytes do not spontaneously undergo apoptosis, suggesting that their rate of hydrogen peroxide production is not involved in this process.


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Table 1. Rates of hydrogen peroxide production by osteoclasts

 
Role of oxidative/nitrative modifications on cytochrome c release and on its activity in apoptosis. Because the rate of hydrogen peroxide production in osteoclasts was found to be high, it could be expected that this or other ROS could exert some oxidative modification on cytochrome c that might result in facilitating its release from mitochondria. Therefore, more information regarding the primary structure of released cytochrome c was required. To this end, we analyzed tryptic digests of cytosolic cytochrome c (which covered 80% of the protein sequence, excluding 8–9 amino acids from the NH2 terminus and from residue 14 to 25, where the heme is covalently bound to the protein) by performing matrix-assisted laser desorption ionization with time-of-flight (MALDI-ToF) and tandem mass spectrometry of selected fragments and found it identical to native mouse cytochrome c, indicating that the primary structure of cytochrome c released from apoptotic cells is not chemically modified differently from the native or original one before the apoptosis event starts (Table 2).


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Table 2. Masses of tryptic peptides of cytochrome c detected with MALDI-TOF mass spectrometry

 
These results were confirmed independently by investigating whether released and bound cytochrome c exhibited any modification as a result of nitrative stress. Nitration of tyrosine residues can have profound influence on cytochrome c physicochemical properties, because Tyr67 residue lies adjacent to the heme pocket where the electron transfer takes place (8, 52) (Fig. 3). MALDI-ToF experiments on tryptic fragments from both bound and released cytochrome c indicated the presence of several peptides that included Tyr67 (peptides 14 through 17), whose masses were increased by 45.9929 as expected for the addition of a nitro group to Tyr67 and/or an oxidized Met (Table 2). No peptide spanning from residues 57 to 74 was observed without nitration. In contrast, fragments that included Tyr97 were obtained in either the unmodified or nitrated forms (compare peptides 4 and 7 with peptide 12). Tyr48 (peptide 9) and Tyr74 (peptide 11) did not appear to be nitrated, because no peptides were obtained or detected with a mass increased by the addition of a nitro group.



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Fig. 3. Three-dimensional (3-D) structure of cytochrome c, indicating the potential target sites for nitration, i.e., Tyr67 and Tyr97.

 
Because at least two Tyr in cytochrome c were nitrated, we investigated whether the release of nitrated cytochrome c was required to initiate, or was associated with, an increased apoptosis. To ascertain whether this association was specific for apoptosis of resting osteoclasts or for other apoptosis-induced processes, we compared the level of nitration of cytochrome c released from control and cycloheximide-treated osteoclasts. We selected cycloheximide treatment as a signaling pathway-independent stimulus to rapidly accelerate osteoclast apoptosis (Fig. 4 and Table 3). Figure 4A shows a 90-min culture of vehicle-treated osteoclasts and documents the presence of both surviving (knobs) and apoptotic osteoclasts (arrows). Figure 4B shows a culture of osteoclasts cultured for 90 min with cycloheximide. It is clear that all of the osteoclasts are apoptotic in the cycloheximide-treated cultures on the basis of distinctive nuclear condensation (arrows). The nitration of cytochrome c was evaluated by probing Western blots for cytochrome c (to identify the band corresponding to this hemoprotein) and reprobing the blots with an antibody specific for 3-nitrotyrosine (Table 3). At 60 min, the apoptosis was three times higher in cycloheximide-treated than in control osteoclasts. This increase in apoptosis was accompanied by a twofold increase in the level of cytochrome c released, whereas the level of nitrated cytochrome c released (by control and cycloheximide-treated osteoclasts) was not significantly different (Table 3). These results indicated that nitrated cytochrome c was not required to initiate apoptosis.



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Fig. 4. Cycloheximide increases osteoclast apoptosis. Purified osteoclasts were cultured for 90 min with a 1:1,000 dilution of either vehicle (H2O; A) or cycloheximide (final concentration 5 µM; B). Cells were fixed and stained for chromatin condensation and TRAP activity as detailed in MATERIALS AND METHODS. Arrows indicate apoptotic osteoclasts; circles indicate viable osteoclasts.

 

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Table 3. Release of cytochrome c, release of nitrated cytochrome c, and apoptosis in control and cycloheximide-treated osteoclasts

 
Notably, if the ratios of nitrated cytochrome c for control and cycloheximide-treated osteoclasts are compared (32 vs. 13% for control and cycloheximide-treated, respectively) with those for apoptosis (22 vs. 66% for control and cycloheximide-treated, respectively), then two observations can be made: 1) less nitrated cytochrome c than native cytochrome c is released from mitochondria; and 2) nitrated cytochrome c is less "active" to trigger apoptosis than native cytochrome c. Regarding the first observation, when experiments were performed to remove all nitrated cytochrome c from osteoclast mitochondria, a higher digitonin-to-protein ratio was required compared with that used for native cytochrome c (i.e., 0.05 mg digitonin/mg protein), indicating that the nitrated species was relatively more tightly bound to mitochondrial membranes, thus requiring higher concentrations of detergent for its full release. Why would nitration of cytochrome c result in a tighter binding to the membrane? To answer this question, we need to review briefly how native cytochrome c is bound to the membrane. Native cytochrome c is bound to the inner mitochondrial membrane by two distinct acidic phospholipid-binding sites (47): a binding provided by ionic interaction [i.e., between conserved Lys at residues 72 and 73 of cytochrome c and negatively charged phosphate groups of cardiolipin (16)] and a conformation provided by hydrogen bonding (i.e., between the amide group of Asn52 and the protonated phosphate group of an acidic phospholipid). In addition, hydrophobic interaction between an expanded acyl chain of cardiolipin and a hydrophobic channel, a channel lined by hydrophobic amino acids leading from the surface of the protein into the heme crevice (10), in cytochrome c anchors the protein to the membrane (45). The ability of this Asn to form hydrogen bonds is evident from studies on the structural water molecules in cytochrome c (39): one water molecule is hydrogen bonded to Asn52, Tyr67, and the carbonyl group of Ile75 (39). This structure is compatible with the close vicinity of one of the cardiolipin phosphates to the heme moiety reported by others (54). Cytochrome c heme is covalently bound to the protein by thioether bonds formed between the vinyl side chains of the heme and two cysteinyl residues of the apoprotein. Two axial coordinate bonds hold the heme in place: a relatively weak bond between the heme iron and one of its axial ligands, the sulfur atom of Met80, and a stronger bond with the other axial ligand, His18 imidazole. These two amino acids form the first tier of protein side chains interacting with the metal. In the second tier of iron-related side chains lies Tyr67, which is located close to the Met80 side chain.

Upon nitration of Tyr67 of cytochrome c with tetranitromethane, several changes were observed that were attributed to various conformational changes as follows. 1) A shift of the 695-nm band heme-linked ionization (the band assigned to the binding of the heme iron by methionyl sulfur; Ref. 49) from a pK of ~9 in the native protein to an apparent pK of 5.9 in the nitrated one has been indicated as being compatible with the increased acidity in 3 pH units of the pK of the phenolic group of Tyr upon nitration (42). 2) Loss of the 695-nm band at neutral pH (50) is indicative of a weakening of the methionyl sulfur and heme iron bond. Thus nitration of Tyr67 residue affects the iron-binding properties of the Met80 sulfur, and accordingly, these two residues might be close to each other in the native molecule, as can be observed in the X-ray crystallography structure of cytochrome c. 3) An increased reactivity of ferricytochrome c with imidazole compared with the native cytochrome c indicates that there is a significant loosening of the closed crevice structure.

Thus, at the relatively acidic pH of the mitochondrial intermembrane space, nitro-Tyr67 will likely undergo intramolecular hydrogen bonding (as in the case of other o-nitrophenols), thus eliminating one of the three hydrogen bonds required to hold the water molecule in the protein interior, liberating Asn52 and Ile75 (and probably Thr58, which is hydrogen bonded to Tyr67) side chains from their unfavorable internal positions. In the particular case of Asn52, as it becomes more exposed, it will favor its interaction with cardiolipin, enhancing the anchorage of cytochrome c to the membrane, and the hydrogen bonding will be favored by the hydrophobic environment (i.e., low dielectric constant) of the membrane (Fig. 5). Our results support this view, because less nitrated cytochrome c than native cytochrome c was found in the cytosol of cells undergoing apoptosis, and higher concentrations of detergent were required to fully separate nitrated cytochrome c from mitochondrial membranes.



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Fig. 5. Model showing the effect of nitration on the release of cytochrome c from the outer leaf of the mitochondrial inner membrane. Left: 3-D structure of cytochrome c indicating residues 52, 75, and 67, the residues involved in the hydrogen bonding of water. When Tyr67 is nitrated and forms an intramolecular hydrogen bond, Asn52 is pushed outward, favoring its interaction with the cardiolipin phosphate heads. Right: upon receiving a certain stimulus, cytochrome c is released from the inner membrane. If the hemoprotein is nitrated, the release is less favorable because of the tighter interaction with cardiolipin.

 
With regard to the second alternative, that nitrated cytochrome c is less "active" to trigger apoptosis than the native one, our results supported this concept (compare ratios of nitrated/native cytochrome c in the cytosol with the apoptosis percentages; Table 3), confirming that native cytochrome c is the active species. Similar conclusions were obtained by other groups: nitrated cytochrome c (obtained by tetranitromethane treatment) was six to seven times less efficient than the native one in restoring maximal respiration of cytochrome c-depleted mitochondria (52) or in sustaining state 4 respiration in intact heart mitochondria (8); increased peroxidatic activity (6) and partial reduction with ascorbate (8, 23) were also observed. However, nitration of cytochrome c did not affect its ability to promote the cleavage of procaspase 3 when mixed with HeLa cytosol extracts, indicating that nitration of this hemoprotein does not suppress apoptosis (40). This apparent discrepancy might reflect the fact that nitrated cytochrome c is not fully inactivated by nitration, requiring relatively large concentrations to restore a particular function, as has been reported for mitochondrial respiration (52).

Other studies were aimed at elucidating the origin of the nitration of cytochrome c in osteoclasts. Because the nitration of biomolecules has been demonstrated to occur as a consequence of a nitric oxide exposure (not necessarily indicating nitric oxide as the actual nitrating species), we investigated whether these cells were endowed with a nitric oxide synthase (NOS). Western blots of osteoclasts (from 10 to 100 µg protein/lane) immunoprobed with monoclonal antibodies to eNOS, nNOS, or iNOS (in the presence of positive controls, to assure the operativity of the technique, negative controls, which included the abrogation of primary antibody addition, and loading controls, probing for actin to assure equal protein loading) were negative, indicating that this enzyme was not present in our osteoclasts that were cultured on plastic (Fig. 6). This finding is in contrast to the studies by Kasten et al. (24). These prior studies examined osteoclasts that were generated in vivo and isolated from bone for study, whereas the present studies examined osteoclasts that were generated in vitro and cultured on plastic. Thus the state of activation of these different populations is a likely explanation for these discrepancies.



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Fig. 6. Western blots of nitric oxide synthase (NOS) in osteoclasts. Western blotting of NOS was performed as described in MATERIALS AND METHODS. Duplicate aliquots of osteoclasts were loaded, as well as a positive control, purchased from the manufacturer: rat pituitary lysate for neuronal NOS (nNOS), mouse macrophage lysate for inducible NOS (iNOS), and human endothelial cell homogenate for endothelial NOS (eNOS). Negative controls included the omission of primary antibodies, and loading controls were performed by reprobing the blots for actin.

 
Role of Bcl-2 family in osteoclast apoptosis. Bcl-2 family members play a critical role in modulating the participation of mitochondria in driving apoptosis (28, 40). Increased osteoclast numbers and size are associated with Paget's disease, and Brandwood et al. (4) have documented that Bcl-2 levels are elevated in Pagetic osteoclasts, showing that altered osteoclast survival is likely to be involved in pathological bone loss. To further study the role of osteoclast mitochondria in apoptosis, we examined cultured purified osteoclasts to determine expression of both prosurvival and proapoptotic members of the Bcl-2 family. Initially, we examined expression levels for several members of this family (Fig. 7A). We observed relatively high expression levels of prosurvival bcl-2 and bcl-x. To determine the impacts of culture of mature osteoclasts on these proteins, we probed Western blots of purified osteoclasts that had been cultured for up to 60 min. As shown in Fig. 7B, the prosurvival family members Bcl-xL and Bcl-2 disappeared before the onset of apoptosis. The apparent disparity in the expression levels at time 0 may simply reflect the different sources and titers of the antibodies used in these blots. These data supported the possibility that the loss of prosurvival Bcl-2 family members might be critical to driving osteoclast apoptosis. Bcl-2 and Bcl-xL have distinctly different as well as shared functions (35, 37). To determine whether these two proteins have similar or different roles in blocking osteoclast apoptosis, we examined whether an increase in Bcl-2 expression in isolated osteoclasts could replace the degraded Bcl-2 and Bcl-xL and promote osteoclast survival. To examine this, we infected mature osteoclasts with an adenoviral expression vector containing wild-type Bcl-2 (Fig. 7, C and D). To confirm that we were successful in increasing osteoclast Bcl-2 protein expression, we examined Western blots of osteoclasts infected with either empty adenoviral vector or a Bcl-2 adenoviral expression vector (Fig 7C). There was detectable Bcl-2 in freshly isolated osteoclasts that decreased with culture when the cells were infected with empty vector (Fig. 7C, 24V). In contrast, infection with Bcl-2 for 24 h (Fig. 7C, 24B) resulted in increased Bcl-2 protein levels, and the protein level increased further with 48 h of infection (Fig. 7C; see 48B). For the impact of infection on osteoclast apoptosis, we examined osteoclasts infected with MOIs estimated at 1 or 8 per nucleus (Fig. 7D). Again, cultures were maintained for either 24 or 48 h before fixing and apoptosis analysis. As shown, there was some impact of vector infection in the first 24 h postinfection (Fig. 7D, 24V), but this was not evident after 48 h of infection (Fig. 7D, 48V). The transient survival supporting influences of vector treatment was surprising. It has been reported that both adenoviral and baculoviral infections can alter cell survival by stimulating TNF-{alpha} and IL-1{alpha} expression (12, 2122, 2829, 51). Because these cytokines also promote osteoclast survival, this may be the cause of the temporary survival enhancement that we observed. At both 24 h (Fig. 7D, 24B) and 48 h (Fig. 7D, 48B) after Bcl-2 infection, osteoclasts infected with a MOI of 8 per nucleus survived better than those infected with a MOI of 1 per nucleus, which survived better than those that were not infected (Fig. 7D, time 0).



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Fig. 7. Bcl-2 family members and osteoclast survival. A: osteoclasts were differentiated, and bcl-2 and bcl-x mRNA levels were assessed using the Mouse Q Series apoptosis gene array. B: mature osteoclasts were purified and either harvested (time 0) or cultured for the indicated time period. Protein expression levels for Bcl-2, Bcl-xL, or tubulin were determined using Western blot analysis. C and D: isolated osteoclasts were infected with either empty vector (V) or Bcl-2 expression vector (B) for either 24 (24V and 24B) or 48 h (48V and 48B) as indicated. C: infection at a MOI of 8 per nucleus. Bcl-2 and tubulin protein expression was determined using Western blot analysis. D: infection at a MOI of 1 and 8 per nucleus. Osteoclast apoptosis was detected as described in MATERIALS AND METHODS. *P < 0.05 compared with no infection (MOI of 0).

 
Role of caspase cascade in osteoclast apoptosis. Many proapoptotic stimuli require a mitochondria-dependent step, controlled by pro- and antiapoptotic members of the Bcl-2 family, leading to cytosolic release of mitochondria intermembrane space proteins that can trigger either caspase activation, such as cytochrome c, or caspase-independent death pathways, such as apoptosis-inducing factor (AIF) (30, 62). Bax is one of the main proapoptotic Bcl-2 family proteins; the presence of either Bax or Bak is required for most mitochondria-dependent cell death processes, including those induced by tBid (the caspase-activated form of the "BH3-domain only" proapoptotic Bcl-2 family protein Bid) and by proapoptotic drugs such as staurosporine and actinomycin D (65). Actinomycin D, similarly to staurosporine, requires the presence of Bax or Bak to induce cytochrome c release and cell death (64). However, treatment of osteoclasts with actinomycin D did not result in a significant change in the number of apoptotic cells compared with either controls or vehicle-treated cells (actinomycin D: 62 ± 4, control: 48 ± 12, vehicle: 58 ± 6). These results suggest either that the release of cytochrome c was already occurring at a maximum rate (thus addition of either of these compounds could not increase the apoptosis any further) or that a step downstream of cytochrome c release was acting as the rate-limiting step. In this regard, it has been reported that activation of given caspases after mitochondrial outer membrane permeabilization may be the limiting step for AIF (a flavoprotein that can stimulate a caspase-independent cell death pathway) detachment from the outer leaf of the inner membrane and cytosolic release (64).

Interactions between certain death receptors such as TNF-R1, Fas, Trail-R1, and Trail-R2 with their ligands leads to the recruitment and autoactivation of initiator procaspase-8 (25). Recent studies indicated that caspase-8 substrates are located at distinct cellular loci, playing key roles in mediating death receptor-induced apoptosis. For example, caspase-8 cleavage of Bid promotes cytochrome c and Smac/Diablo release from mitochondria (31, 68). More recently, caspase-8 cleavage of BAP31 at the endoplasmic reticulum (ER) resulted in the formation of p20, which mediates Ca2+-dependent apoptotic cross talk between ER and mitochondria, stimulating mitochondrial fission and sensitization of these organelles to caspase-8-induced cytochrome c release (5). The importance of caspase-8 cleavage in our experimental model (and that of those pathways downstream from caspase-8) was tested by incubating osteoclasts with 2 µM z-IETD-fmk, an inhibitor of caspase-8. Treatment of osteoclasts with the inhibitor for 90 min resulted in 60 ± 2% apoptosis, a moderate decrease compared with that of vehicle-treated cells (73 ± 3%). The lack of a significant inhibition of apoptosis upon the inhibition of caspase-8 supports two conclusions. First, the release of cytochrome c is not restricted to caspase-8 as an effector caspase. Indeed, caspases other than caspase-8 (i.e., caspase-3, caspase-6, and caspase-7) have been shown to trigger rapid release of cytochrome c (3). Second, the p20 signaling pathway may not be a requirement for cytochrome c release in this experimental model but rather a sensitizer of this event, as suggested by others (3).

To investigate whether the aforementioned caspases are active participants in osteoclast apoptosis, we studied the role of both initiator (i.e., upstream regulators) and effector caspases (i.e., degraders of structural proteins and cellular protective mechanisms) in osteoclasts by using various specific inhibitors. Our results indicated that the initiator caspase-4, caspase-5, and caspase-9 were most important in promoting apoptosis and that they likely targeted activation of effector caspase-3-like caspases (caspase-3 and caspase-7) (Fig. 8). The effectiveness of this relatively low inhibitor concentration could be due to the low osteoclast density once support cells are removed (Fig. 1). Experimental evidence for the cleavage (activation) of caspase-3 and caspase-9 was provided by Western blotting of osteoclasts at 0 and 90 min of culture (Fig. 9). Partial caspase cleavage was detectable as early as 30 min (data not shown). These data confirm that initiator caspase-9 and effecter caspase-3 are involved in osteoclast apoptosis.



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Fig. 8. Role of caspases in osteoclast apoptosis. The role of caspases in osteoclast apoptosis was investigated using 2 µM of each inhibitor or vehicle. The inhibitors z-LEHD-fmk and z-DEVD-fmk (Chemicon) target caspase-4, -5, and -9 and caspase-3 and -7, respectively. The percentage of apoptotic cells was evaluated.

 


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Fig. 9. Activation of caspase-3 and -9. The activation of caspase-3 and -9 and the level of tubulin was followed by Western blotting with the use of specific antibodies to these proteins (Cell Signaling).

 
ROS and apoptosis. Because it was clear that oxidative/nitrative stress did not introduce modifications in cytochrome c to facilitate its release and/or activity in apoptosis, further experiments were performed to ascertain the role of hydrogen peroxide production by mitochondria in apoptosis. To this end, the rate of hydrogen peroxide production by purified, cultured osteoclasts was evaluated at various time points. This rate increased linearly during the first 10 min of culture, reaching a plateau at 30 min and maintaining this rate until the last time point was evaluated (i.e., 90 min). Given that ROS (i.e., hydrogen peroxide, superoxide anion, and hydroxyl radical) produced as a consequence of oxidative stress might initiate apoptosis and considering that the onset of the rate of hydrogen peroxide production was an early event, we investigated the role of these species in osteoclast apoptosis. To this end, the mitochondrial rate of hydrogen peroxide production was modulated in osteoclasts by adding FCCP and/or spin traps. FCCP is an uncoupler of electron transport and oxidative phosphorylation that decreases the rate of hydrogen peroxide production by mitochondria, and spin traps [DMPO or {alpha}-(4-pyridil-1-oxide)-N-tert-butylnitrone (POBN)] were used at concentrations required to trap superoxide anion and hydroxyl radicals. The hydrogen peroxide concentration and the apoptosis of osteoclasts treated with these compounds were evaluated at 60 min of incubation (Table 4). The concentration of hydrogen peroxide in FCCP-treated cells was 10 times lower than that present in control cells, and apoptosis was 2.2 times lower (Table 4). The apoptosis of these cells followed a reciprocal association with the concentration of hydrogen peroxide, indicating a prominent role for mitochondria-derived ROS in apoptosis. Interestingly, hydrogen peroxide was also shown to increase osteoclast differentiation in a chicken marrow-derived cell line (55). The possibility that hydrogen peroxide could support differentiation of precursors and induce apoptosis of mature cells is not unlike the multiple roles of TNF-{alpha} and hematopoiesis as TNF-{alpha} supports survival of some differentiation stages while driving apoptosis in other stages (1, 11 20, 58, 63). Our results indicated that hydrogen peroxide, produced by mitochondria, had a critical role in modulating osteoclast apoptosis; however, it was not clear whether hydrogen peroxide per se was the direct activator of apoptosis or whether hydrogen peroxide served as the precursor of a free radical derived from it, possibly hydroxyl radical.


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Table 4. Effect of spin traps on concentration of hydrogen peroxide and on osteoclasts' apoptosis

 
To investigate whether hydroxyl radical, derived from the cleavage of hydrogen peroxide, was an initiator of apoptosis, we designed experiments to test the effect of spin traps, namely, DMPO (and POBN; not shown) on apoptosis. If an oxygen free radical were involved in the apoptotic process, then the addition of a spin trap, which combines with a free radical, should halt the process at the step mediated by the radical. Addition of DMPO to osteoclasts resulted in an almost 50% decrease in apoptosis of both control and FCCP-treated cells (Table 4). Analysis of the EPR spectra under these conditions allowed the identification of the free radicals involved in the process. The EPR spectrum of control cells resulted in a composite of two major species, superoxide anion (detected as DMPO-OOH adduct; aN = 14.3 G and aH= 11.7) and hydroxyl radical (detected as DMPO-OH adduct; aN = aH = 14.8 G) in a 3:1 ratio (Fig. 10). The EPR signal intensity (which is proportional to the amount of free radicals present, evaluated by double integration of both signals) was 4.5 times lower (on average) when FCCP was present, consistent with an intensity 4 times lower when the effects of decreasing hydrogen peroxide by FCCP addition (2 times lower) and adding the spin trap (2 times lower) are combined.



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Fig. 10. Electron paramagnetic resonance (EPR) spectra of spin trap adducts obtained from control and FCCP-treated osteoclasts. Osteoclasts were treated with 200 mM 5,5'-dimethyl-1-pyrrolyne-N-oxide (DMPO) without (control) or with 1 µM FCCP. EPR spectra were recorded at 60 min of incubation in a Varian E-102 spectrometer. The assignments to each free radical ({bullet}, superoxide anion; {circ}, hydroxyl radical) were performed by using published hyperfine splitting constants for the aforementioned radicals and simulating the signals using the simulation package from NIH. Instrumental settings are described in MATERIALS AND METHODS.

 
A mechanism consistent with these observations would entail primarily the formation of superoxide anion by mitochondria, dismutation of this species to hydrogen peroxide, and production of hydroxyl radical, formed during the splitting of the O-O bond of hydrogen peroxide. These conclusions are supported by the effect of FCCP, a mitochondrial uncoupler, which significantly decreased the concentration of hydrogen peroxide (Table 4) and decreased the EPR signal of superoxide anion and hydroxyl radical (Fig. 10). It could be surmised that the protective role of FCCP could be achieved by preventing the production of ATP by mitochondria, thus depleting the system of ATP, a molecule required to promote apoptosis. However, if this were the case, cell viability would have been compromised as a result of the lack of ATP, required to sustain all ATP-dependent processes. This apparent controversy can be bridged by considering glycolysis as the main source of ATP despite the high mitochondrial density of these cells. This is in agreement with studies from the '70s until recently in which 80–90% of the ATP levels in cultured cells has been demonstrated to be provided by glycolysis (41, 62) without the active participation of the tricarboxylic acid cycle.

In conclusion, excessive bone loss is a major pathology in several diseases, including periodontitis, postmenopausal osteoporosis, glucocorticoid-induced osteoporosis, and metastatic tumor-driven osteolysis. These diverse diseases share a common denominator in the elevation in the numbers of osteoclasts present during bone degradation. Because osteoclast numbers are controlled by factors that have an impact on the rates of differentiation and elimination by apoptosis, understanding the mechanisms by which osteoclasts die may be important in future therapeutic designs to limit the number of osteoclasts.

Our results indicated that the mitochondrial apoptotic pathway is active in osteoclasts, as has been shown for other cell types. The sequence of events upon removal of stromal cells seems to be an increase in the steady-state concentration of hydrogen peroxide, which leads to a higher formation of hydroxyl radical, release of cytochrome c from mitochondria, formation of apoptosome in the cytosol, and activation of caspase-3-like and caspase-9. It is clear that native, unmodified cytochrome c was released by mitochondria to amplify apoptosis and that this hemoprotein did not have a different compartmentalization, as has been proposed by others, or significant oxidative/nitrative modifications. The events that lead to the increase in hydrogen peroxide, release of cytochrome c, and other processes that may contribute to osteoclast apoptosis are currently under investigation in our laboratories. However, we can hypothesize either that stromal cells contribute to the maintenance of the steady-state concentration of hydrogen peroxide in osteoclasts (given that this compound is a freely diffusible species) or that the removal of these cells results in a decrease in macrophage colony-stimulating factor and receptor activator of NF-{kappa}B ligand (RANKL), which may alter the coordinated regulation of the transcriptional program during the in vitro differentiation of osteoclasts. Among the RANKL-induced subset of signaling molecules, there are many protein kinases, docking proteins, and small G proteins and their regulators. Within the expression profile for genes preferentially induced by RANKL, CDC2-related protein was shown to be upregulated in response to RANKL at day 1, decreasing steadily by day 6 (7). It was reported (9) that purified peroxiredoxins I, II, III, and IV (a family of enzymes capable of catabolizing hydrogen peroxide, in which II and V are present in mitochondria) could be phosphorylated on Thr90 by cyclin-dependent kinases (CDKs), including CDC2 (also known as CDK1), resulting in a marked inhibition of the peroxidase activity of this enzyme. In this report (9), it was shown that CDC2, the CDK that is activated during mitosis, is responsible for peroxiredoxin phosphorylation in intact cells (HeLa, HepG2, and NIH/3T3) and that accumulation of hydrogen peroxide that results from the inactivation of peroxiredoxin might be important for progression of the cell cycle. In our experimental model, the decrease in RANKL could lead to an upregulation of CDC2, resulting in the inactivation of peroxiredoxin through CDC2-mediated phosphorylation. The increase in hydrogen peroxide would then result in an increased hydroxyl radical production, which by increasing the permeabilization of the outer mitochondrial membrane or by other mechanisms not yet known, would favor the release of cytochrome c and activation of the apoptotic pathway. Future studies are needed to focus on understanding the chemical characteristics of these steps that lead to osteoclast apoptosis.


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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS AND DISCUSSION
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This work was supported by the Department of the Army Grant DAMD17-00-1-0346, the Minnesota Medical Foundation, and the Lilly Center for Women's Health (to M. J. Oursler) and in part by National Institutes of Health Grants GM-66768 and ES-011407 (to C. Giulivi) and DE-14680 (to M. J. Oursler).


    ACKNOWLEDGMENTS
 
We thank Genevieve Gorny, Virginia Haynes, and Laura Yager for excellent technical support. Digital imaging and processing of gels and blots were performed at the Visualization and Digital Imaging Laboratory, University of Minnesota, Duluth.

Present address of M. J. Oursler: Endocrine Research Unit, Mayo College of Medicine, Rochester, MN 55905 (E-mail: Oursler.MerryJo{at}mayo.edu).


    FOOTNOTES
 

Address for reprint requests and other correspondence: C. Giulivi, Dept. of Molecular Biosciences, School of Veterinary Medicine, University of California, Davis, 1095 Haring Hall, One Shields Ave., Davis, CA 95616-8741 (E-mail: cgiulivi{at}ucdavis.edu)

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


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