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Keywords:

  • apoptosis;
  • ATP;
  • cerebellar granule cells;
  • cytochrome c release;
  • mitochondria;
  • necrosis

Abstract

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

In rat cerebellar granule cells, cytochrome c release takes place during glutamate toxicity and apoptosis due to deprivation of depolarising levels of potassium. We show that, as in necrosis, the released cytochrome c present in the cytosolic fraction obtained from cerebellar granule cells undergoing apoptosis can operate as a reactive oxygen species (ROS) scavenger and as a respiratory substrate. The capability of the cytosolic fraction containing cytochrome c, obtained from cerebellar granule cells undergoing either necrosis or apoptosis, to energise coupled mitochondria isolated by the same cells is also investigated. We show that, in both cases, the cytosolic fraction containing cytochrome c, added to mitochondria, can cause proton ejection, and membrane potential generation and can drive ATP synthesis and export in the extramitochondrial phase, as photometrically measured via the ATP detecting system.

Cytochrome c, separated immunologically from the cytosolic fraction of apoptotic cells when added to mitochondria, is found to cause proton ejection to generate membrane potential and to drive ATP synthesis and export in a manner not sensitive to the further addition of the cytosolic fraction depleted of cytochrome c, which failed to do this. In the light of these findings we propose that in apoptosis the released cytochrome c can contribute to provide ATP required for the cell programmed death to occur.

Abbreviations used
BME

basal medium Eagle

CCCF

cytosolic fraction containing cytochrome c

CF

cytosolic fraction

CGCs

cerebellar granule cells

C-GLU-cells

control glutamate-treated cells

CN

cyanide

cyt

c, cytochrome c

DIV

days in vitro

e.u.

enzymatic units

FCCP

carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone

GDH

glutamate dehydrogenase

GLU-cells

glutamate-treated cells

GNT

glutamate neurotoxicity

mAb-cyt

c, cytochrome c bound to the monoclonal antibody

PBS

phosphate buffer saline medium

RAM

rotenone, anti-mycin and myxothiazole

ROS

reactive oxygen species

S-K25

cells, control cells

S-K5

cells, apoptotic cells

SOD

superoxide dismutase

In neurones, as well as in many other cell types, death can take place either according to a programmed route generally referred to as apoptosis, or as a result of toxic insults of various types and origin. The mechanism(s) through which the two types of death occur could take advantage of the availability of cell model systems undergoing either necrosis or apoptosis according to the stimuli. In vitro cultured rat cerebellar granule cells (CGCs) encounter a necrotic type of death when exposed to excessive and prolonged glutamate exposure (Calissano et al. 1993) and via apoptosis when deprived of depolarising levels (25 mm) of extracellular potassium (D'Mello et al. 1993; Choi 1994).

We have recently reported an apoptosis/necrosis transition in these neurones when the cellular anti-oxidant system is pharmacologically impaired, pointing to reactive oxygen species (ROS) as a major player of such a transition (Atlante et al. 2003). ROS are also responsible for the release of mitochondrial cytochrome c (cyt c) (Atlante et al. 2001; Bobba et al. 2002) which participate in caspase activation and execution of apoptosis (Li et al. 1997; Bossy-Wetzel et al. 1998). Thus, although a causal correlation between the amount and timing of ROS production on one side and the release of cyt c on the other side is widely documented (Atlante et al. 2000), little is known about their possible and actual interplay in both necrotic and apoptotic pathways, especially in connection with the role of mitochondria and ATP production.

Since in necrosis released cyt c can work as ROS scavenger and respiratory substrate (Atlante et al. 2000), in this paper we investigate whether such activity is detectable also in CGCs undergoing apoptosis and, more importantly, whether the released cyt c once reduced can generate the electrochemical proton gradient and drive ATP synthesis and export outside mitochondria in the same cultured neurones undergoing both types of cell death. We also investigate whether in apoptosis the released cyt c can work either per se or together with other molecules present in the cytosol and/or released from mitochondria.

Reagents

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Tissue culture medium and fetal calf serum (FCS) were purchased from Gibco (Grand Island, NY, USA) and tissue culture dishes were from NUNC (Taastrup, Denmark). All enzymes and biochemicals were from Sigma Chemicals Co. (St Louis, MO, USA). Monoclonal anti-cytochrome c antibodies against either the denatured protein (7H8–2C12) or against the native protein (6H2.B4) were purchased from Pharmingen (San Diego, CA, USA); anti-glutamate dehydrogenase antibodies were kindly supplied by Dr F. Rothe (Institut fuer Medizinische Neurobiologie, University of Magdeburg, Germany). Protein A/G PLUS-agarose were purchased from Santa Cruz Biotechnology (Santa Cruz, California, USA).

Cell cultures

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Primary cultures of CGCs were obtained from dissociated cerebellar of 7-day-old Wistar rats as in Levi et al. (1984). Cells were plated in basal medium Eagle (BME) supplemented with 10% FCS, 25 mm KCl, 2 mm glutamine and 100 µg/mL gentamicin on dishes coated with poly-l-lysine. Cells were plated at 2 × 106 per 35 mm dish or 6 × 106 per 60 mm dish. 1β-Arabinofuranosylcytosine (10 µm) was added to the culture medium 18–22 h after plating to prevent proliferation of non-neuronal cells.

Apoptosis induction

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Apoptosis was induced as in D'Mello et al. (1993): at 6–7 days in vitro (DIV), cells were washed twice and switched to a serum-free BME (S-), containing 5 mm KCl and supplemented with 2 mm glutamine and 100 µg/mL gentamicin for the time reported in the figure legends. Apoptotic cells are referred to as S-K5 cells. Control cells were treated identically but maintained in serum-free BME medium supplemented with 25 mm KCl for the indicated times and are then referred to as S-K25 cells.

Glutamate neurotoxicity induction

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Glutamate exposure was performed 7 days after plating. Primary cultures were exposed for 30 min to glutamate (100 µm) at 25°C in Locke's solution (154 mm NaCl, 5.6 mm KCl, 3.6 mm NaHCO3, 2.3 mm CaCl2, 5.6 mm glucose, 10 mm Hepes pH 7.4) in the presence of 1 µm glycine added in order to fully activate NMDA-sensitive glutamate recognition sites (Johnson and Ascher 1987). Cells were then replenished with BME containing 25 mm KCl, 2 mm glutamine and gentamicin (100 µg/mL) and put in the incubator. For the quantitative assessment of glutamate neurotoxicity (GNT), cell integrity and count were measured, as described in (Volontèet al. 1994), after 12–24 h. Glutamate-treated cells and control cells are referred as GLU- and C-GLU-cells, respectively.

Assessment of neuronal viability

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Viable CGCs were quantified by counting the number of intact nuclei after dissolving the cells in detergent-containing solution as described in Volontèet al. (1994). Apoptosis or glutamate neurotoxicity was expressed as the percentage of intact cells with respect to control cells kept under the same respective experimental conditions. In control experiments 95–97% integrity was found after 24 h.

DNA fragmentation analysis

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Fragmentation of DNA was performed as in Hockenbery et al. (1990). Briefly CGCs (6 × 106) were plated in poly l-lysine-coated 60-mm tissue culture dishes, collected with cold phosphate-buffered saline (PBS, pH 7.2); after removal of the medium and washing once with cold PBS, CGCs were centrifuged at 3500 g for 5 min. The pellet was lysed in 10 mm Tris-HCl, 10 mm EDTA, 0.2% Triton X-100 (pH 7.5). After 30 min on ice, the lysates were centrifuged at 17 000 g for 10 min at 4°C. The supernatant was digested with proteinase K and then extracted twice with phenol-chloroform/isoamylic alcohol (24 : 1). The aqueous phase containing soluble DNA was recovered, and nucleic acids were precipitated with sodium acetate and ethanol overnight. The pellet was washed with 70% ethanol, air-dried and dissolved in TE buffer (10 mm Tris-HCl, 1 mm EDTA, pH 7.5). After digestion with RNase A (50 ng/mL at 37°C for 30 min), the sample was subjected to electrophoresis in a 1.8% agarose gel and visualised by ethidium bromide staining. Soluble DNA from equal numbers of cells was loaded in each lane.

Cell homogenate, mitochondria and cytosolic fraction preparation

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Before each experiment, the culture medium was removed and the plated CGCs, i.e. S-K5- or GLU-cells and the respective control cells were washed with PBS containing 138 mm NaCl, 2.7 mm KCl, 8 mm Na2HPO4, 15 mm KH2PO4 pH 7.4, and then collected by gentle scraping in a final volume of 4 mL PBS/90-mm dish. Suspended granule cells showed full viability, even though they lacked the morphological organisation present in culture dishes such as cell–cell and cell–substrate contacts as well as neuritis. Cell integrity, which remains constant for 3–5 h, was quantitatively assessed by checking the inability of cells to oxidise externally added succinate, which cannot enter intact cells (Berry et al. 1991), by checking the ability of ouabain to block glucose transport in cells (Sjodin 1989), and by counting dead cells, identified as large phase-bright cell bodies, as in Volontèet al. (1994). The final cell suspension routinely contained 85–95% intact cells and was prepared after 6–7 DIV.

Cell homogenate from a cell suspension was obtained by 10 strokes with a Dounce potter at room temperature. With this procedure lactate dehydrogenase is released and subsequent treatment with Triton-X-100 does not cause further release.

Mitochondria were isolated from either S-K5- or GLU-CGCs and their respective control, essentially as reported in Almeida and Medina (1998). Briefly, the cell homogenate was centrifuged at 1500 g for 10 min at 4°C and the supernatant was kept on ice. The pellet was re-homogenized with a further 3 mL of isolation buffer, consisting of 320 mm sucrose, 1 mm K+-EDTA, 10 mm Tris-HCl pH 7.4, and the homogenate was centrifuged at 1500 g for 5 min at 4°C. The two supernatants were pooled and centrifuged at 1500 g for 10 min at 4°C. Supernatant was further centrifuged at 17 000 g for 11 min at 4°C. The pellet, i.e. the mitochondrial fraction, was then re-suspended in 200 µL of isolation buffer to obtain about 4 mg of mitochondrial protein/mL. Mitochondria were shown to be essentially pure without significant cytosolic contamination as shown by checking a variety of cytosolic marker enzymes, such as lactate dehydrogenase (E.C.1.1.1.27), citrate synthase (E.C.4.1.3.7) and acetylcholinesterase (E.C.3.1.1.7), (Almeida and Medina 1998) in the mitochondrial preparation. On the other hand, these mitochondria, incubated in PBS, were checked for their integrity and coupling by measuring the activities of both adenylate kinase (ADK, E.C.2.7.4.3) and glutamate dehydrogenase (GDH, E.C.1.4.1.3) (see below), which are marker enzymes of the mitochondrial intermembrane space and matrix, respectively. The percentage of damaged mitochondria ranged between 0.5 and 1.5%. Mitochondrial coupling was checked by measuring the respiratory control ratio (RCR), i.e. (oxygen uptake rate after ADP addition)/(oxygen uptake rate before ADP addition) which reflects the ability of mitochondria to produce ATP; in both cases succinate was used as a respiratory substrate. As expected, both the inhibitors of electron flow and atractyloside, a powerful inhibitor of the ADP/ATP translocator (LaNoue and Schoolwerth 1984), blocked ADP-stimulated oxygen uptake. Inhibition was also caused by oligomycin which can inhibit ATP synthase (Nicholls 1982) and rapidly reversed by the uncoupler FCCP, which stimulates the rate of oxygen consumption.

Cytosolic fractions of both control and S-K5 or GLU-CGCs (about 30 × 106cells in 1 mL) were obtained after homogenization in PBS and centrifugation at 15 000 g for 15 min.

The supernatant was concentrated until to 0.2 mL, by using SAVANT Speed Vac Concentrator, and indicated as cytosolic fraction containing cyt c (CCCF). Cytosolic fractions obtained from S-K5- and GLU-CGCs are referred as CCCFA and CCCFN, respectively.

Both the protein assay of cell and mitochondria were determined according to Waddel and Hill (1956), with bovine serum albumin used as a standard.

Polarographic measurements

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

O2 consumption was measured polarographically by means of a Gilson 5/6 oxygraph (Gilson Medical Electronics Inc., Middletown, WI, USA) using a Clark electrode, as in (Atlante et al. 1996, 2000). Instrument sensitivity was set to a value in order to follow rates of O2 uptake as low as 0.5 natoms min−1mg−1 protein. The cell homogenate in PBS (about 0.2 mg protein) was incubated in a thermostated (25°C) water-jacketed glass vessel (final volume equal to 1.5 mL).

In order to detect polarografically the cyt c presence in the cytosolic fraction (CCCF), the capability of CCCFN/A to oxidise ascorbate was checked. Because ascorbate cannot permeate the outer mitochondrial membrane (Alexandre and Lenhinger 1984) its oxidation can occur as a result of the release from mitochondria of a component that can oxidise ascorbate and then reduce oxygen via cytochrome c oxidase, i.e. cyt c present in the cytosolic fraction (Atlante et al. 2001).

Safranine 0 response assay

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Safranine 0 response was monitored as in Passarella et al. (1990). Time-dependent absorbance changes were recorded with a Jasco double-beam/double-wavelength spectrophotometer (Jasco Corporation, Tokyo, Japan) UV-550 wavelengths of 520 and 554 nm. Measurements were carried out at 25°C in 2 mL of standard medium consisting of 200 mm sucrose, 10 mm KCl, 1 mm MgCl2, 20 mm HEPES-Tris pH 7.2, containing 1 µm safranine 0 and 0.1 mg protein.

Proton movement measurements

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Proton movement across the mitochondrial membrane was followed, as previously reported (Passarella et al. 1990; de Bari et al. 2002) by means of a Gilson 5/6 Oxygraph, equipped with a Gilson pH 5 Servo Channel electrode (Gilson Medical Electronics Inc., Middletown, WI, USA) which allows for continuous monitoring and direct recording of mitochondrial suspension pH changes during the assays. Mitochondria (0.1 mg protein) were added, at 25°C, to 1.5 mL of the proton medium consisting of 100 mm NaCl, 10 mm MgCl2, 1 mm EGTA-TRIS, 2 mm TRIS-HCl, pH 7.0.

ADP/ATP carrier measurement

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Mitochondria from CGCs (0.1 mg protein) were incubated at 25°C in 2 mL of standard medium. The ATP appearance in the extramitochondrial phase as a result of externally added ADP, was revealed by using the ATP detecting system (ATP D.S) consisting of glucose (2.5 mm), hexokinase (HK, E.C.2.7.1.1) (0.5 e.u.), glucose-6-phosphate dehydrogenase (G6PDH, E.C.1.1.1.49) (0.5 e.u.) and NADP+ (0.2 mm) (Passarella et al. 1988). The rate of NADP+ reduction in the extramitochondrial phase was observed photometrically as the absorbance increased to 340 nm, measured as a tangent to the initial part of the progress curve and expressed as nmoles NADP+ reduced/min × mg mitochondrial protein. The ε340 value for NADH was found to be 6.4 mm/cm.

Immunodepletion of cytochrome c from cytosolic fraction

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

Monoclonal anti-cytochrome c antibody (6H2.B4) that recognises the native form of cyt c was used. An aliquot of 20 µL of this antibody (0.5 mg/mL IgG 1) was incubated with the same volume of Protein A/G PLUS-agarose beads re-suspended in 40 µL of PBS at 4°C for 3 h. The beads were collected by centrifugation for 15 min at 1000 g. After the removal of the supernatant, the beads were washed once with 1 mL of buffer, consisting of 320 mm sucrose, 1 mm K+-EDTA, 10 mm Tris-HCl pH 7.4, and incubated with 2.5 mL CCCFA (1 mg proteins) for 5 h at 4°C. The beads were subsequently pelleted by centrifugation for 15 min at 1000 g, the resulting supernatant is the cytosolic fraction immunodepleted of cyt c (CFA) and the pellet is the cyt c bound to the anti-cytochrome c antibody (mAb-cyt c). In both cases the cytochrome c presence was checked as described above.

Apoptosis/necrosis evaluation in CGCs

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

In order to establish criteria useful to identify how granule cerebellar cells die, they were compared against each other with respect to the cell survival time course, and to the occurrence of DNA laddering (Fig. 1). No differences between CGCs subjected to glutamate exposure (100 µm; 30 min) (Calissano et al. 1993) and potassium shift (D'Mello et al. 1993) were found with respect to cell survival up to 8 h; later death occurred faster via necrosis than apoptosis, with no survival found at 24 h when about 50% of cells in which apoptosis was induced still were live (Fig. 1a). In agreement with D'Mello et al. (1993) CGCs apoptosis was inhibited by either 1 µg/mL actinomycin D or 10 µg/mL cycloheximide, inhibitors of transcription and translation, respectively, that were ineffective in rescuing CGCs undergoing necrotic death. On the other hand, glutamate neurotoxicity, but not apoptosis, was fully prevented by (+/–)-5-methyl-10,11-dihydro-5-H-dibenzo(a,d)cyclohepten-5,10-imine hydrogen maleate (MK-801, 1 µm), a non-competitive antagonist of the NMDA receptor (not shown). DNA laddering was found only for cells en route of apoptosis after 8–15 h (Fig. 1b, lanes d–e), in this regard, the necrotic cells are indistinguishable from the control (Fig. 1b, compare lanes g–h and I–j).

image

Figure 1. Cell survival and DNA fragmentation in CGCs undergoing apoptosis or necrosis. (a) Time course of loss of viability of CGCs undergoing apoptosis (S-K5) or necrosis (GLU). Cells were either switched to serum-free medium containing low K+ (S-K5) for different times or treated with glutamate (100 µm for 30 min) (see Materials and methods) for the same times (GLU). Cell viability was determined by counting the number of intact nuclei and is expressed as the percentage of S-K25- or C-GLU-cells to which a 100% value was given. Control values were 100 ± 5. Results are means ± standard errors of triplicate measurements and representative of six different experiments carried out with different cell preparations from different groups of animals. (b) DNA fragmentation. Soluble DNA was extracted from either neurones switched to serum-free culture medium containing low K+ (S-K5) for 8 and 15 h (lanes d and e) or neurones treated with glutamate for the same times (lanes I and j). Lanes (b and c) and (g, and h) contain DNA from either control cells maintained in high K+ (S-K25) or C-GLU-cells (c) for 8 h and 15 h, respectively. DNA from equal numbers of plated cells (6 × 106) was loaded in each lane. Size marker was HaeIII-digested ΦX174 phage DNA (lanes a, f, and k).

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In each experiment of this work we have investigated cell culture plates from the same preparation with respect to the above reported criteria, thus confirming on an experimental basis that the cell used at the times indicated, i.e. 30 min and 3 h of glutamate exposure and potassium shift, respectively, will die via necrosis and apoptosis.

The fraction containing the released cyt c can operate as a ROS scavenger and respiratory substrate in mitochondria isolated from CGCs undergoing apoptosis.

A set of experiments was carried out in order to gain some insights into the role played by the cyt c released in the extramitochondrial phase during apoptosis. First the presence of cytochrome c in the cytosolic fraction (CCCFA), obtained from S-K5-CGCs at different times from apoptosis induction, was checked by western blot using a monoclonal anti-cytochrome c antibody (mAb-cyt c) that recognises the denatured form of the protein (Fig. 2a). Polyclonal antibodies against glutamate dehydrogenase (GDH) were used to normalize the corresponding quantity of cyt c revealed on the same filter. The cyt c proved to be already detectable in the cytosolic fraction 10 min after apoptosis induction in increasing quantities up to 3 h, but declining thereafter, as in Bobba et al. (1999). As expected (Atlante et al. 2003), the presence of vitamin E and C and SOD, separately added, was found to strongly reduce the quantity of cyt c in the extramitochondrial phase (not shown).

image

Figure 2. The fraction containing the released cyt c can work as a ROS scavenger and a respiratory substrate in cells undergoing apoptosis. (a) Cytosolic fractions (CCCFA) prepared from either S-K25- or S-K5-CGCs with different treatment times were analysed by immunoblotting with a monoclonal antibody that recognises the denatured cyt c. Antibodies against GDH were used to normalise the amount of protein loaded onto the gel (for further details see Materials and methods). (b) CCCFA (200 µL) (prepared as described in Materials and methods), from either Control or S-K5-CGCs (see Materials and methods) with different treatment times, was incubated in 1.5 mL of PBS in the presence of xanthine oxidase (1 e.u./mL). Cyt c reduction was started with 10 µm xanthine (XX) addition and measured as absorbance decrease at 550 nm. Variations of up to 5% were found in five different experiments, carried out with different cell cultures prepared from different groups of animals. (c) Mitochondria (0.2 mg protein), isolated from rat CGCs cultured for 7 DIV are incubated in a polarographic vessel at 25°C in 1.5 mL of PBS. At the arrows, the following additions were made: 10 µm xanthine (XX) plus 1 e.u./mL xanthine oxidase (XOD), 200 µL CCCF from either control or S-K5-CGCs at different treatment times, 1 mm cyanide (CN), 10 µm allopurinol (ALLO). The rate of oxygen uptake is expressed as nanoatoms O2/min × mg mitochondrial protein. Variations of up to 5% were found in five different experiments, carried out with different cell cultures prepared from different groups of animals. The insets b′ and c′ describe the processes occurring (see text). Either the oxidised or the reduced form of CCCF are referred as CCCFOX and CCCFRED, respectively. COX = cytochrome c oxidase.

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CCCFA was checked for its capability to work as an electron acceptor from ROS, i.e. as a ROS scavenger, and as a respiratory substrate as previously reported in CGCs under glutamate neurotoxicity (Atlante et al. 2000).

The capability of the CCCFA to oxidise superoxide anion to oxygen was analysed by monitoring cyt c reduction photometrically as an increase of absorbance at 550 nm (Fig. 2b). In order to check the existence of oxidised cyt c in the extract, cyt c reduction via dithionite (Errede et al. 1978) was verified (not shown). CCCFA preparations, obtained by CGCs undergoing apoptosis for times ranging from 10 min to 8 h, were added to the ROS producing system, consisting of 10 µm xanthine plus 1 e.u./mL xanthine oxidase (XX + XOD). Consistently with Fig. 2(a), as the quantity of reduced cyt c increased with increasing times from the apoptotic trigger up to 3 h. For longer times, a decrease in the reduced cyt c was found as expected as the quantity of cyt c diminishes as a result of the caspase-dependent proteolysis (Bobba et al. 1999). As a control, we checked that no cyt c reduction occurred in the cytosolic fraction obtained from both S-K25 and from S-K5 cells incubated in the presence of SOD (50 U/106 cells), used as a ROS scavenger (not shown).

In a parallel assay, carried out with the same cells, DNA laddering was found to occur essentially as in Fig. 1(b). The same assay was currently carried out in this work under similar conditions and it will be not mentioned later.

To ascertain whether cyt c reduced as a result of superoxide anion formation can function as an electron donor to molecular oxygen, mitochondria were added with XX plus XOD (Fig. 2c) and oxygen consumption was measured in the absence (control) or presence of CCCFA preparations. In controls, i.e. in the absence of CCCFA or where the added cytosolic fraction is obtained from S-K25 cells, oxygen consumption starts when XX + XOD is added with a cyanide-independent rate equal to 12 nanoatoms O2/min × mg cell protein and it is strongly inhibited by allopurinol, the specific XOD inhibitor (Flower et al. 1985; Atlante et al. 1997). When CCCFA obtained from CGCs undergoing apoptosis for times ranging from 10 min to 8 h were present, the rate of oxygen consumption was progressively higher up to 3 h after the apoptotic trigger and declined thereafter, which is in agreement with the data shown in Fig. 2(a) and (b). Externally added cyanide (CN) was found to decrease the rate of oxygen uptake compared with that obtained in the presence of ROS producing system either alone or plus the cytosolic fraction obtained from control cells (control). No oxygen consumption was found following the addition of allopurinol. These findings suggest that the (XX + XOD)-dependent oxygen consumption is dependent both on the O2 consumption due to superoxide formation (Fig. 2, b′, c′) and on the level of cyt c released, which is reduced by ROS and, in turn, it reduces molecular oxygen to form water via cyt c oxidase. Thus, as far as CCCFA is concerned, the findings reported in Fig. 2 are consistent with a two-phase process characterized first by the reduction of cyt c present in the cytosolic fraction (Fig. 2b′), followed by the oxidation of the reduced cyt c acting as an electron donor for cyt c oxidase (COX) (Fig. 2c′). The decrease in the oxygen uptake found for CCCFA obtained from cells undergoing apoptosis for 5–8 h is consistent with the caspase-dependent cyt c proteolysis (Bobba et al. 1999; Atlante et al. 2003).

The cytosolic fractions containing the released cytochrome c can cause protonmotive force generation in mitochondria isolated from either GLU-CGCs or S-K5-CGCs.

The capability of cytosolic fractions, obtained from either CGCs exposed to glutamate, i.e. undergoing necrosis (CCCFN), or from sister cultures undergoing apoptosis (CCCFA), to cause both proton ejection from the mitochondrial matrix and membrane potential generation was checked by directly measuring proton concentration change outside mitochondria and by measuring safranine absorbance changes as Δψ probe.

In the case of glutamate neurotoxicity, cytosolic fractions were obtained from cells incubated for 30 min with 100 µm glutamate (see Materials and methods), when mitochondria are still coupled and about 40% cyt c release occurs (Atlante et al. 1999, 2000). As a control both the negligible survival at 24 h and the lack of DNA laddering was found, as in Fig. 1, with CGCs from the same preparation. Apoptotic fractions, on the other side, were obtained after 3 h when mitochondria are essentially coupled and maximum cyt c release occurs (Atlante et al. 1998; Bobba et al. 1999).

The presence of cyt c in both CCCFN and CCCFA was ascertained by western blot analysis (Fig. 3a) with GDH used to normalise the released cyt c, as described above.

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Figure 3. The capability of the cytosolic fraction containing the released cyt c to cause protonmotive force generation in mitochondria isolated from either GLU-CGCs or S-K5-CGCs. (a) Cytosolic fractions contain the released cyt c. Cytosolic fractions (CCCFN/A) prepared from either C-GLU- and GLU-CGCs (30 min glutamate-treatment, necrosis) or S-K25- and S-K5-CGCs (3 h K+ shift, apoptosis), were analysed by immublotting with a monoclonal antibody that recognises the denatured cyt c. Antibodies against GDH were used to normalise the amount of protein loaded onto the gel (for further details see Materials and methods). (b) Proton movements. Mitochondria (0.1 mg protein), isolated from either GLU-CGCs (30 min glutamate-treatment, necrosis) or S-K5-CGCs (3 h K+ shift, apoptosis), were suspended at 25°C in 1.5 mL of proton medium consisting of 100 mm NaCl, 10 mm MgCl2, 1 mm EGTA-TRIS, 5 mm TRIS-HCl (pH 7.00) in the presence of rotenone (3 µm), anti-mycin (0.8 µm) and myxothiazole (6 µm). At the arrows the following addition were made: ascorbate (ASC, 5 mm), CCCFN/A (200 µL) (prepared as described in Materials and methods) and FCCP (1.25 µm). Proton movement was monitored as described in Materials and Methods. (c) Safranine response. Mitochondria (0.1 mg protein), isolated from either GLU-CGCs (30 min glutamate-treatment, necrosis) or S-K5-CGCs (3 h K+ shift, apoptosis), were suspended at 25°C in 2 mL of PBS containing safranine (1 µm) in the presence of rotenone (3 µm), anti-mycin (0.8 µm) and myxothiazole (6 µm). When the absorbance trace remained constant the following additions were made: ascorbate (ASC, 5 mm), CCCFN/A (200 µL) (prepared as described in Materials and methods) and FCCP (1.25 µm). Safranine response was monitored as described in the Materials and methods. (d) Polarographic measurement. Mitochondria (0.1 mg protein), isolated from either GLU-CGCs (30 min glutamate-treatment, necrosis) or S-K5-CGCs (3 h K+ shift, apoptosis), were suspended at 25°C in 2 mL of PBS and oxygen amount was measured as a function of time. At the arrows the following additions were made: ascorbate (ASC, 5 mm), CCCFN/A (200 µL) (prepared as described in the Materials and methods), ADP (1 mm) and cyanide (CN, 1 mm).

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No change in the proton concentration outside mitochondria was found in the presence of 5 mm ascorbate, used as reducing agent, and rotenone, anti-mycin and myxothiazole (RAM), used to inhibit complex I–III of the respiratory chain. As shown in Fig. 3(b) addition of CCCFN/A caused proton ejection from mitochondria isolated from either glutamate-treated or S-K5 cultures. As expected, proton uptake by mitochondria was found following the addition of the uncoupler FCCP.

When mitochondria were isolated from cells undergoing either necrosis or apoptosis and tested under the above conditions, Δψ generation following the CCCFN/A addition was detected as revealed by the decrease of safranine absorbance (Fig. 3c). Δψ generation was found to be prevented by cyanide (not shown) and abolished by FCCP. In the same experiment, either succinate (5 mm) or pyruvate (0.5 mm), used as a control (not shown), generated membrane potential with a higher rate with respect to CCCFN/A in a manner inhibited by anti-mycin and by rotenone, respectively.

To check that isolated mitochondria with CCCFN/A added could take up O2 in a manner controlled by ADP, mitochondria were isolated from either glutamate-treated or by K+-deprived CGCs, pre-incubated with RAM plus ascorbate, and CCCFN/A were added to the mixture. Measurements of oxygen uptake made in both State 4 and State 3 in the presence of 1 mm ADP (Fig. 3d) demonstrate an oxygen uptake with a control respiratory index equal to 2.1 and 2.5, respectively. Cyanide completely blocks oxygen consumption. In the same experiment, oxygen uptake was caused by either the substrate pair glutamate plus malate, completely inhibited by rotenone, or by succinate, the oxidation of which was completely blocked by anti-mycin A, a powerful inhibitor of complex III; oxygen consumption was restored by adding ascorbate plus TMPD substrate pair, which directly reduced cyt c, but completely inhibited by cyanide (not shown).

The cytosolic fractions containing the released cytochrome c can drive ATP synthesis and export in mitochondria isolated from either GLU-CGCs or S-K5-CGCs.

The finding that externally added CCCFN/A can cause oxygen uptake stimulated by ADP implies that it is endowed with the capability to generate the protonmotive force (Fig. 3). In order to ascertain that mitochondria are still able to carry out oxidative phosphorylation and that CCCFN/A, once reduced, can drive ATP synthesis and make the newly synthesised ATP available for extramitochondrial reactions, we resorted to a procedure (Passarella et al. 1988) that allowed for the continuous monitoring of ATP efflux from mitochondria with ADP added (Fig. 4). In a typical experiment mitochondria from control CGCs (control), and with AP5A added, used to inhibit adenilate kinase (Lienhard and Secemski 1973), were incubated in the presence of the ATP D.S. (see Materials and methods). The ATP concentration outside mitochondria is negligible as no increase in the absorbance measured at 340 nm is found in the presence of glucose, hexokinase, glucose-6-phosphate dehydrogenase and NADP+. As a result of ADP (0.1 mm) addition, an increase in the NADPH absorbance was found, at a rate equal to 12.5 nmoles NADP+ reduced/min × mg mitochondrial protein, indicating the appearance of ATP in the extramitochondrial phase (Fig. 4a). Strong inhibition in the rate of absorbance increase occurs in the presence of either atractyloside or carboxyatractyloside, inhibitors of the ADP/ATP translocators, or oligomycin, the inhibitor of ATP synthase (not shown). When mitochondria were added with the ROS producing system, i.e. XX + XOD, the rate of ATP production was about 5 nmoles/min × mg mitochondrial protein with a strong inhibition (60%) likely as a result of the ROS formation (see Atlante et al. 1989). No ATP production was found in the mitochondria added with RAM. On the contrary, XX + XOD addition to mitochondria, added with RAM and incubated with CCCFN/A, which fails to change the measured absorbance, proved to restore ATP production at a rate equal to 2.5 and 3.2 nmoles/min × mg mitochondrial protein for necrosis and apoptosis conditions, respectively (Fig. 4a).

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Figure 4. CCCFN/A drives ATP synthesis and export from mitochondria of either GLU-CGCs or S-K5-CGCs. Mitochondria (0.1 mg protein), isolated from control CGCs (control) (a), GLU-CGCs (30 min glutamate-treatment, necrosis) (b) or S-K5-CGCs (3 h K+ shift, apoptosis) (c), were incubated at 25°C in 2.0 mL of standard medium consisting of 0.2 m sucrose, 10 mm KCl, 1 mm MgCl2, 20 mm HEPES-Tris pH 7.2 in the presence of 5 mm ascorbate [only in (b) and (c)], 10 µm Ap5A, and ATP D.S. (see Materials and methods). At the arrows ADP (100 µm) is added. When present, RAM, in the absence or presence of either xanthine (10 µm) plus xanthine oxidase (1 e.u./mL) (XX + XOD) [in (a)] or CN [in (b) and (c)], and CCCFN/A (200 µL) (prepared as described in Materials and methods) were added 2 min and 1 min before ADP addition, respectively. The ATP appearance was measured as described in the Materials and methods. Numbers along the traces give the rate of NADP+ reduction measured as the tangent at the initial part of progress curve and expressed as nmoles NADP+ reduced/min × mg protein.

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In light of results that show inhibition as a result of ROS, in other experiments, ascorbate was used to reduce cyt c both in necrosis and apoptosis, as a result of ADP addition to mitochondria. ATP efflux in the extramitochondrial phase was measured with a rate equal to 6.8 and 7.4 nmoles/min × mg mitochondrial protein (Figs 4b and c), showing the same inhibitor sensitivity of the control. When electron flow in complex I and III and then ΔpH and Δψ generation were prevented by previously adding mitochondria with RAM, no ATP appearance was detected (Figs 4b and c) both in the presence or absence of externally added ascorbate. Addition of the CCCFN/A, obtained from CGCs treated with glutamate for 30 min and from S-K5-CGCs after 3 h of potassium shift, caused an ATP efflux from mitochondria at rates equal to 3.1 and 4.9 nmoles/min × mg mitochondrial protein, respectively. Such an effect was completely prevented by CN (Figs 4b and c).

The released cyt c, but not the cytosolic fraction depleted of cyt c, can cause protonmotive force generation and drive ATP synthesis and export in mitochondria isolated from apoptotic neurones.

During apoptosis, but not during glutamate neurotoxicity, a variety of proteins/factors, besides cyt c, is released from mitochondria (Susin et al. 1999; Adrain et al. 2001; van Loo et al. 2001; Cande et al. 2002). In order to determine whether the mitochondria energisation, reported in Figures 3 and 4, is a result of the combined action of different proteins or of cyt c alone, use was made of the monoclonal anti-cyt c antibody (6H2.B4), which recognises the native protein, to remove cyt c from the CCCFA (see Materials and methods). The complete removal of cyt c from CCCFA was shown by the lack of immunoreactivity in the fraction (CFA) obtained from the antibody-treated CCCFA in a western blot analysis (Fig. 5a). Then, the two fractions obtained, i.e. the cyt c-depleted cytosolic fraction (CFA) and the cyt c bound to the antibody (mAb-cyt c), were checked with respect to their capability to energize mitochondria. As shown in Fig. 5, no proton movement and Δψ generation were found when CFA was added to mitochondria incubated with RAM plus ascorbate, whereas the addition of the mAb-cyt c fraction caused both proton ejection outside mitochondria and Δψ generation, which were reversed by externally added FCCP (Figs 5b and c). In a parallel experiment, both proton ejection and Δψ generation were found as a result of the addition of mAb-cyt c only to the mitochondria (Figs 5b′,c′). The rates of both proton ejection and Δψ generation measured in b–b′ and c–c′ were found to be over imposable in three distinct experiments carried out with mitochondria and cellular extracts derived from different cultures.

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Figure 5. Ab-cyt c generates ΔµH+ and works as a respiratory substrate in cells undergoing apoptosis. (a) Cyt c-depleted cytosolic fraction. Both CCCF and the cytosolic fraction immunodepleted of cyt c (CFA) were analysed by immunoblotting with a monoclonal antibody that recognises the denatured protein. Antibodies against GDH were used to normalise the amount of protein loaded onto the gel. (b) Proton movements. Mitochondria (0.1 mg protein), isolated from S-K5-CGCs (3 h K+ shift, apoptosis), were suspended at 25°C in 1.5 mL of proton medium consisting of 100 mm NaCl, 10 mm MgCl2, 1 mm EGTA-Tris, 5 mm Tris-HCl (pH 7.00) in the presence of rotenone (3 µm), anti-mycin (0.8 µm) and myxothiazole (6 µm). At the arrows the following additions were made: ascorbate (ASC, 5 mm), CFA (200 µL) (prepared as described in Materials and methods) and Ab-cyt-c (final concentration about 0.35 µm), prepared from S-K5-CGCs (3 h K+ shift), and FCCP (1.25 µm). Proton movement was monitored as described in the Materials and methods. (c) Safranine response. Mitochondria (0.1 mg protein), isolated from S-K5-CGCs (3 h K+ shift, apoptosis), were suspended at 25°C in 2 mL of PBS containing safranine (1 µm) in the presence of rotenone (3 µm), anti-mycin (0.8 µm) and myxothiazole (6 µm). When the absorbance trace remained constant the following additions were made: ascorbate (ASC, 5 mm), CFA (200 µL) (prepared as described in Materials and methods) and Ab-cyt c (final concentration about 0.35 µm), prepared from S-K5-CGCs (3 h K+ shift), and FCCP (1.25 µm). Safranine response was monitored as described in the Materials and methods. (d) Polarographic measurement. Mitochondria (0.1 mg protein), isolated from S-K5-CGCs (3 h K+ shift, apoptosis), were suspended at 25°C in 2 mL of PBS and oxygen amount was measured as a function of time. At the arrows the following additions were made: ascorbate (ASC, 5 mm), CFA (200 µL) (prepared as described in Materials and methods) and Ab-cyt c (final concentration about 0.35 µm), prepared from S-K5-CGCs (3 h K+ shift), ADP (1 mm) and cyanide (CN, 1 mm).

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In the same experiments ADP-stimulated oxygen uptake, which reflects the oxidative phosphorylation, was monitored (Fig. 5d-d′). No oxygen consumption occurs when CFA was added to mitochondria, but these organelles uptake oxygen when added with mAb-cyt c in an ADP-stimulated manner. It is worth noting that no difference in oxygen consumption rate was found if mAb-cyt c is added in the presence or absence of CFA (compare Figs 5 d,d′).

In another set of experiments, the capability of both CFA and mAb-cyt c in driving ATP synthesis and efflux from mitochondria isolated from apoptotic cells was compared (Fig. 6) by using the ATP detecting system (ATP D.S). Externally added ADP (0.1 mm) was proven to cause ATP appearance outside mitochondria isolated from S-K5-CGCs and added with Ap5A (not shown). When ADP was added to S-K5 mitochondria incubated with RAM, no ATP efflux was measured (Fig. 6a), further CFA addition failed to cause ATP appearance in extramitochondrial phase, whereas as a result of the mAb-cyt c addition, a fast NADP+ reduction was found to occur at a rate of about 5 nmoles/min × mg protein. A similar rate was measured when CFA was added before ADP (Fig. 6b). In Figs 6(c) and (d), ATP efflux was measured as started by the ADP addition to mitochondria previously added with mAb-cyt c in the absence or presence of CFA. The rates of ATP appearance outside mitochondria were similar to those in 6(a) and 6(b). Therefore, the results reported in Fig. 6 show that mAb-cyt c provides the energy needed for ATP synthesis and ADP/ATP exchange to occur.

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Figure 6. mAb-cyt c drives ATP synthesis and export from mitochondria isolated from S-K5-CGCs. Mitochondria (0.1 mg protein), isolated from S-K5-CGCs (3 h K+ shift, apoptosis), were incubated at 25°C in 2.0 mL of standard medium consisting of 0.2 m sucrose, 10 mm KCl, 1 mm MgCl2, 20 mm HEPES-Tris pH 7.2 in the presence of 10 µm Ap5A, 3 µm rotenone, 0.8 µm anti-mycin, 6 µm myxothiazole, 5 mm ascorbate and ATP D.S. (see Materials and methods). At the arrows the following additions were made: ADP (100 µm), CFA (200 µL) (prepared as described in Materials and methods) and mAb-cyt c (final concentration about 0.35 µm), prepared from S-K5-CGCs (3 h K+ shift). Cyanide (CN), when present, is added together with RAM. The ATP appearance was measured as described in Materials and methods. Numbers along the traces give the rate of NADP+ reduction measured as tangent at initial part of progress curve and expressed as nmoles NADP+ reduced/min × mg protein.

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Discussion

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

One of the most outstanding problems in bioenergetics concerns the role of mitochondria in the processes leading to cell death. Their major role in the glutamate-dependent necrosis is commonly accepted, dying neurones have been shown to lose their mitochondrial membrane potential and energy charge (Ankarcrona et al. 1995), and a key role in Ca2+ homeostasis (Budd and Nicholls 1996; Schinder et al. 1996; Nicholls and Budd 1998) was proposed for mitochondria that were also shown to undergo Ca2+-dependent uncoupling (Dugan et al. 1995). An early and progressive mitochondrial dysfunction (Atlante et al. 1996; Schinder et al. 1996) and oxidative stress (Coyle and Puttfarcken 1993; Lafon-Cazal et al. 1993; Atlante et al. 1997) have been shown to occur under GNT (Atlante et al. 2001). Consistently, an increase of glucose uptake by CGCs was shown to account for the higher rate of anaerobic glycolysis (Minervini et al. 1997).

We have recently reported that in glutamate-dependent death, cyt c is released from mitochondria, and that it can operate as an ROS scavenger and as a respiratory substrate (Atlante et al. 2001). Cyt c release also occurs during apoptosis (Du et al. 1997; Kluck et al. 1997a; Bossy-Wetzel et al. 1998; Jürgensmejer et al. 1998; Bobba et al. 1999, 2002; Gorman et al. 1999; Ott et al. 2002; Scorrano et al. 2002) and it has been proposed that it plays a role in caspase activation in an ATP-dependent process (Li et al. 1997; Ueda et al. 1998). How cyt c is released from mitochondria remains to be elucidated: it was hypothesised that the cyt c release occurs with the involvement of the proteins forming the permeability transition pore, either with or without voltage-dependent anion channel (VDAC) (Kantrow and Piantadosi 1997; Petit et al. 1998; Shimizu et al. 1999). However, it is worth noting that the bidirectional cyt c movement across the outer mitochondrial membrane has been shown to occur without adenylate kinase release (Atlante et al. 1999; Wieckowski et al. 2001) from intact coupled mitochondria, i.e. in which the integrity of the inner membrane is maintained, before a loss of mitochondrial Δψ (Atlante et al. 2001; Bossy-Wetzel et al. 1998 and Waterhouse et al. 2001). Finally, cyt c release proved to be insensitive to permeability transition pore inhibitors (Bossy-Wetzel et al. 1998; Neame et al. 1998; Krohn et al. 1999; Goldstein et al. 2000; Wigdal et al. 2002).

Apart from the manner by which cyt c is released from mitochondria, the question then arises as to the functional role played by the cytoplasmic pool of cyt c relationship with necrotic or apoptotic death. We show that, as in glutamate neurotoxicity, in CGCs undergoing apoptosis the fraction containing the cyt c, released from intact and respiring mitochondria, can transfer electrons from the superoxide anion to molecular oxygen via the respiratory chain (Fig. 2), i.e. it functions as an ROS scavenger and as a respiratory substrate. The inhibition of oxygen uptake caused by CN demonstrates that O2 reduction takes place via cyt c oxidase, thus we conclude that the cyt c present in the CCCFA can be oxidised via this oxidase. We further demonstrated that by transferring electrons to the cytochrome oxidase in mitochondria, in which any oxidation of endogenous substrate is prevented, the released cyt c can generate the electrochemical proton gradient and drive ATP synthesis. The fact that cyt c-dependent oxygen uptake provides proton ejection and Δψ generation is shown here (Fig. 3) by using mitochondria in which ATP synthesis is monitored by the increase of the rate of oxygen consumption caused by ADP addition. Moreover, in the presence of cytosolic fractions, ATP synthesis and export from mitochondria isolated from CGGs undergoing necrosis or apoptosis (Fig. 4) occurs in spite of the presence of inhibitors, which prevent energy formation as a result of the oxidation of endogenous substrates.

The direct oxidation of cyt c by cytochrome oxidase at mitochondrial contact sites (Ardail et al. 1990) seems unlikely in view of oxidase structure and apparent inability to span the two mitochondrial membranes. Moreover, a structurally distinct type of contact sites, named bridge contact sites, was revealed in brain mitochondria (Perkins et al. 2001), which might play a role in maintaining the structural integrity of the outer and inner membrane systems, as well as in allowing cyt c oxidation. In addition, the cyt c bound to its antibody (mAb-cyt c) is also recognised by mitochondria and oxidised by cytochrome c oxidase, this is consistent with the observation that the monoclonal anti-cyt c antibody (6H2.B4) binds to a region around residue 62 of rat cytochrome c, which in the three-dimensional structure of mammalian cyt c is located on the back of the molecule relative to the heme (Goshorn et al. 1991).

In the light of these findings, we suggest that in CGC apoptosis cyt c is released from mitochondria as a holoenzyme, and that the redox state in which it sits is not constant and can therefore be affected by the presence of redox substances, including ROS which are abundantly generated in vivo both in glutamate-triggered necrosis and during apoptosis.

This leads to the conclusion that the redox state of the cyt c affects its function in the ATP production and, consequently, in the activation of caspase (Li et al. 1997; Ueda et al. 1998) finally allowing for the progression of apoptosis. Consistently, in our experiments, ascorbate was strictly required to reduce cyt c and then energise mitochondria. It should be noted that, in another cell system, the cyt c released from mitochondria was suggested to work as a pro-apoptotic factor independent of its redox state (Kluck et al. 1997b; Hampton et al. 1998).

Comparisons made between this paper and Atlante et al. (2000) show that the amount of the released cyt c from cell undergoing apoptosis or necrosis is quite similar at the respective times considered for the experiments, i.e. 3 h for apoptosis and 30 min for necrosis, but in the light of both the differences existing between the two death types, and the experimental manipulations required in the mitochondria preparations, no comparison with respect to the released cyt c efficiency can be made.

However, at least in the early phase of both necrosis and apoptosis, cyt c release plays a major role but with opposite features. In necrosis cyt c release and the related changes in cell metabolism, namely NADH oxidation via mitochondrial NADH-b5 oxidoreductase as well as plasma membrane NADH oxidoreductase and lactate production, are evoked to counterbalance the deficit in glucose oxidation (Atlante et al. 1999) and then contribute to the maintenance of adequate cell ATP, but cell damage is almost complete in 5 h of glutamate treatment. Contrarily, in agreement with the ATP and cyt c requirement for the induction of the apoptotic program (Liu et al. 1996; Nicotera et al. 2000), we show that the released cyt c plays a crucial role in channeling the reducing equivalent, perhaps ROS, in energy source for apoptotis to continue. Thus, while the cyt c release with the consequent ATP production plays a role in the cell defence to antagonise the fast and massive insult evoked by glutamate, the same processes can participate in the progression of the apoptotic pathway from the commitment to the execution phase.

As far as other roles of the released cyt c are concerned, it is worth mentioning that upon entering the cytosol, cyt c cooperates with at least two other factors in the proteolytic activation of the specific caspases (Liu et al. 1996; Kluck et al. 1997b; Zou et al. 1997).

Indeed, it should be noted that mitochondria contain and release proteins, besides cyt c, involved in the apoptotic cascade (Susin et al. 1999; van Loo et al. 2001; Cande et al. 2002), such as AIF (apoptosis inducing factor), which seems to induce nuclear condensation (Hu et al. 1998; Patterson et al. 2000; Zamzami and Kroemer 2001) and a protein that promotes certain forms of apoptosis, possibly by neutralising one or more members of the family of anti-apoptotic proteins, i.e. Smac/DIABLO (Du et al. 2000; Adrain et al. 2001; Ekert et al. 2001). However, because the cytosolic fraction depleted of cyt c fails in causing proton ejection and Δψ generation, and consequently to drive ATP synthesis (Figs 4 and 5), we are forced to conclude that, at least in CGCs, the pool of cytosolic and mitochondrial released components do not contain, besides cyt c, any equivalent reducing acceptor to be used in oxidative phosphorylation. Thus, the presence of cytosolic fractions does not influence the pool of mAb-cyt c in its action of ATP production outside mitochondria. We therefore conclude that no other protein released from mitochondria participate in the cyt c-dependent processes reported here.

This paper shows that cyt c acts as a bi-functional factor, it can remove ROS as a scavenger and can drive ATP synthesis, thus regulating the apoptosis time course. Such a bi-functional activity is probably significant during the commitment phase, lasting the first 3–8 h before the proteolytic cascade which, at least in CGCs, takes place in the execution phase which starts after such an interval (Bobba et al. 1999; Canu et al. 2000; Atlante et al. 2003). At present we cannot exclude that in the cytosol the released cyt c acting as an ROS scavenger requires other cytosolic/mitochondrial components, although it appears unquestionable that it is the last component allowing electrochemical proton gradient generation and ATP synthesis. Whether the rate of ATP appearance in the extramitochondrial phase is comparable to that generated by the transport via the ADP/ATP carrier as found elsewhere (Passarella et al. 1988), or to that of the ATP synthase reaction (Gagliardi et al. 1997) cannot be presently established. Interestingly, a major role of the ADP/ATP carrier in the regulation of cell-death (Vieira et al. 2000), via a possible interaction with VDAC (Desagher and Martinou 2000) or due to excessive ROS production (Kantrow et al. 2000) has been proposed in other apoptotic systems. The elucidation of these biochemical events is presently under active investigation.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References

The authors thank Mr Vito Petragallo for his excellent technical assistance. This work was partially financed by PRIN ‘Metabolismo Energetico e Trasporto Mitocondriale nella Morte Cellulare ed in altre Condizioni Fisio-Patologiche’ (MURST), by Fondi di Ricerca di Ateneo del Molise to SP, by Ministero della Salute grant ‘Alzheimer Project’ to PC and by FISR ‘Stress ossidativo e bioenergetica mitocondriale nella patogenesi delle malattie neurodegenerative’ to AA.

References

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Reagents
  5. Cell cultures
  6. Apoptosis induction
  7. Glutamate neurotoxicity induction
  8. Assessment of neuronal viability
  9. DNA fragmentation analysis
  10. Cell homogenate, mitochondria and cytosolic fraction preparation
  11. Polarographic measurements
  12. Safranine 0 response assay
  13. Proton movement measurements
  14. ADP/ATP carrier measurement
  15. Immunoblot analysis
  16. Immunodepletion of cytochrome c from cytosolic fraction
  17. Results
  18. Apoptosis/necrosis evaluation in CGCs
  19. Discussion
  20. Acknowledgements
  21. References
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