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Biological Sciences and Health Center (CCBS), Cruzeiro do Sul University, Regente Feijó Avenue, 1295, Analia Franco, 03342-000 São Paulo, Brazil
1 Department of Physiology and Biophysics, Institute of Biomedical Sciences, University of São Paulo, Avenue Prof. Lineu Prestes, 1524, 05508-900 Butantan, São Paulo, Brazil
(Correspondence should be addressed to R Otton; Email: rosemari.otton{at}unicsul.br)
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Introduction |
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As newly reported by our group, diabetic patients and all oxan-induced diabetic rats presented a high number of apoptotic lymphocytes in blood and mesenteric lymph nodes as shown by the increased number of cells with fragmented DNA, phosphatidylserine externalization, and mitochondria depolarization (Otton et al. 2004). In the same study, we found decreased number of circulating lymphocytes in diabetic patients possibly as a consequence of the high occurrence of apoptosis. Recently, our group also showed that 24- and 48-h fasting was able to cause a considerable increase in the proportion of rat lymphocytes in apoptosis. Concomitantly, plasma levels of FFAs were markedly increased reaching 0.99 mmol/l after 48-h fasting as compared with the control group (0.45 mmol/l; Pires et al. 2007). Treatment of lymphocytes from healthy human subjects as well as leukemia cell lines (Raji and Jurkat) with a fatty acid mixture that mimics the proportion and concentration found in human plasma raises the proportion of cells in apoptosis. Leukemia cells were more susceptible to show apoptosis than lymphocytes from healthy human. These findings support the proposition that conditions which raise plasma fatty acid levels (e.g. diabetes and starvation) may impair immune function by causing lymphocyte death (Otton & Curi 2005).
Fundamental metabolic processes occur in mitochondria, which are essential for the survival of all eukaryotic cells. Apoptosis, however, is also controlled within this organelle. Examples of survival functions are the oxidative phosphorylation (ox-phos) of adenine, electron transport in the respiratory chain, ß-oxidation of fatty acids, as well as the citric acid cycle (trichloro acetic acid (TCA); Mayer & Oberbauer 2003). Mitochondrial ox-phos is the major ATP source in eukaryotes. In this process, electrons released from reducing substrates are delivered to O2 via a chain of respiratory H+ pumps. These pumps (complexes I–IV) establish an H+ gradient across the inner mitochondrial membrane, and the electrochemical energy of this gradient is then used to drive ATP synthesis by the complex V (ATP synthase). Chemically, the stepwise reduction of O2 proceeds via several reactive oxygen species (ROS). These ROS can damage cellular components such as proteins, lipids, and DNA, but recent evidence also highlights a specific role of mitochondrial ROS in the control of cell signaling (Brookes et al. 2004).
ROS and reactive nitrogen species (RNS) can be either harmful or beneficial to living systems (Valko et al. 2007). Beneficial effects of ROS occur at low/moderate concentrations and involve physiological roles in cellular responses to anoxia, as for example, in defense against infectious agents and in the function of a number of cellular signaling systems. The harmful effect of free radicals occurs in biological systems when there is an overproduction of ROS/RNS on one side and a deficiency of enzymatic and non-enzymatic antioxidants on the other (Valko et al. 2001, Ridnour et al. 2005).
A growing body of evidence indicated that oxidative stress, which occurs in cells exposed to elevated FFAs, can cause an increase in cytosolic calcium concentration in addition to a direct effect on mitochondria, causing the mitochondrial permeability transition pore (MPTP) to open irreversibly leading to loss of mitochondrial transmembrane potential, release of cytochrome c, and inhibition of mitochondrial respiration. Calcium has also been linked to the loss of mitochondrial transmembrane potential that accompanies apoptosis. Apparently, the MPTP opens in response to abnormally high levels of calcium that accumulate in mitochondria during apoptosis. On the other hand, mitochondria actively orchestrate the spatiotemporal profiles of intracellular calcium, under both physiological and pathological conditions (Gunter et al. 2000, 2004).
The aim of this study was to investigate the mechanism of human lymphocyte death induced by the same FFAs mixture used in our previous study (Otton & Curi 2005). For this purpose, the following measurements were performed: production of superoxide anion, nitric oxide (NO), intracellular Ca2+ concentration ([Ca2+]i), and mitochondrial cytochrome c release. In addition, we investigated whether the FFA mixture changes the expression of pro- (FasL, bad, and bid) and anti-apoptotic (14-3-3) proteins and the activities of caspases 3, 6, 8, and 9. The experiments were performed in freshly obtained cells and in 24-h cultured lymphocytes.
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All reagents for buffers, fatty acids, Histopaque-1077, HEPES, EGTA, ionomycin, and BSA were obtained from Sigma Chemical Co. RPMI-1640 culture medium, antibiotics, and fetal calf serum were purchased from Invitrogen. Polyclonal anti-cytochrome c, anti-bid, anti-bad, anti-14-3-3, anti-FasL, and anti-
-actin antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Ac-DEVD-AMC (caspase 3), Ac-VEID-AMC (caspase 6), Ac-IETD-AFC (caspase 8), and Ac-LEHD-AMC (caspase 9) were from Pharmingen–BD Biosciences (San Diego, CA, USA). Lucigenin, dihydroethidium, DAF-2-FM, Fura-2-AM, pluronic acid, and propidium iodide were obtained from Molecular Probes (Eugene, OR, USA).
Cell isolation and culture
The Human Ethic Committee of the Institute of Biomedical Sciences and Faculty of Medicine, São Paulo University, approved the research protocol of this study. Peripheral blood lymphocytes obtained from healthy subjects were collected by venopuncture procedure and placed in vacuum/siliconized test tubes containing heparin as anticoagulant agent. Blood samples (20 ml) were diluted in the same volume of 150 mM NaCl solution and lymphocytes were separated using Histopaque 1077 according to the manufacturer’s instructions (Sigma). After centrifugation, peripheral blood mononuclear cells were collected from the interphase and maintained in RPMI-1640 medium for 1 h to allow the adherence of monocytes so as to obtain a pure lymphocyte preparation (about 98%). Afterwards, human lymphocytes were grown in RPMI-1640 medium supplemented with 2 mM glutamine, 20 mM HEPES, 10% fetal calf serum, 10 U/ml penicillin, and 10 µg/ml streptomycin, maintained at 37 °C and 5% CO2 humidified atmosphere for up to 48 h when indicated.
Cell treatment
The fatty acid mixture used in the present study was prepared to mimic the proportion of fatty acids found in normal human plasma (Otton & Curi 2005). The proportion of fatty acids was as follows: 1.74% lauric (C12:0), 5.2% myristic (C14:0), 31% palmitic (C16:0), 1.1% palmitoleic (C16:1), 41% stearic (C18:0), 4.6% oleic (C18:1), 9.6% linoleic (C18:2), 1.3% linolenic (C18:3), 3.2% arachidonic (C20:4), 0.45% eicosapentaenoic (C20:5), and 1.8% docosahexaenoic (C20:6) acids. The concentrations of the fatty acids used varied from 0.1 to 0.4 mM. The fatty acid concentrations used are usually found in plasma of healthy humans (Butler et al. 2001, Bajaj et al. 2002, Clore et al. 2002, Woerle et al. 2002). The percentage of ethanol was always lower than 0.05% of the total volume of the culture medium. This concentration of ethanol has shown not to be toxic for the cells (Siddiqui et al. 2001, Lima et al. 2002). All experiments were performed with cells left untreated (control) or treated with ethanol (vehicle). BSA was added at 0.2% as an extra cellular fatty acid chelator. There was no difference between untreated and ethanol-treated cells in all cases.
Assays of the apoptotic features
Apoptosis was assessed by several criteria. DNA fragmentation was quantified by cell cycle analysis of total DNA content as described by Nicoletti et al.(1998). Quantitative determination and differentiation of viable, early, and late apoptotic cells were carried out using annexin V (20 µg/ml)/propidium iodide (50 µg/ml) staining as previously described (Fadok et al. 1992). The rhodamine 123 assay was carried out to investigate the mitochondrial transmembrane potential (
m) in cells treated with the fatty acid mixture for 48-h culture as described by Johnson et al.(1980). Rhodamine 123 is a membrane-permeable lipophilic cationic fluorochrome that serves as a probe of mitochondrial transmembrane potential (Bedner et al. 1999). Rhodamine 123 accumulates in mitochondria, and the extent of its uptake, indicated by the intensity of cellular fluorescence, reflects the mitochondrial m (Darzynkiewicz et al. 1981, Lizard et al. 1995, Castedo et al. 1996). The attraction of the cationic rhodamine 123 molecules by the relatively high negative electric potential across the mitochondrial membrane may be the basis for the selective staining of this organelle in living cells (Johnson et al. 1980). The specificity of rhodamine 123 as mitochondrial probe is greater when this fluorochrome is used at low concentration (Johnson et al. 1981). The drop in mitochondrial m, which is an important event of apoptosis, is reflected by a decrease in retention (of rhodamine 123 and so the green fluorescence). After culture, cells were harvested, centrifuged, washed with PBS, and incubated with rhodamine 123 at 5 µM for 15 min at 37 °C in the dark. Afterwards, the cells were washed twice with cold PBS and incubated for 30 min at 37 °C in the dark. Fluorescence of rhodamine 123 was determined using the FL1 channel (green fluorescence 530/30 nm) in a flow cytometry (488 nm wavelength excitation). As control, cells treated with rotenone (10–5 M) for 30 min, an inhibitor of mitochondrial electron transport chain, showed a pronounced decrease in green fluorescence. All results represent the average obtained in triplicate samples. The variations among the triplicates were always < 10%. All experiments were repeated thrice.
Production of ROS using lucigenin-enhanced chemiluminescence assay
Lucigenin is extensively used to measure the extracellular superoxide anion content mainly produced through NADPH oxidase activation. After being excited by superoxide anion, lucigenin releases energy in the form of light and the chemiluminescence produced can be monitored by a luminometer. Lucigenin (1 mM) was added to cells incubated (2.5 x 106/ml) in the presence of fatty acid mixture at 0.1, 0.2, 0.3, and 0.4 mM in Tyrode’s buffer (NaCl 137 mM, KCl 2.68 mM, MgCl 0.49 mM, NaHCO3 12 mM, NaH2PO4 0.36 mM, D-glucose 5.6 mM, and acid HEPES 5 mM), pH 7.4. The experiments were carried out in the presence and absence of BSA (0.2%). Diphenylene iodonium (DPI; 10 µM), an inhibitor of NADPH oxidase (Chen et al. 2007), was used to investigate if superoxide anion production occurred through NADPH oxidase activation. The chemiluminescence response was monitored for 30 min at 37 °C, in a microplate luminometer (EG&G Berthold LB96V, Bundoora, Australia).
Measurement of ROS by flow cytometry
Dihydroethidium was used for the flow cytometric measurement of intracellular superoxide content. Dihydroethidium is a lipophilic probe and readily diffuses across cell membranes. Once inside the cell, it is rapidly oxidized to ethidium (a red fluorescent compound) by superoxide (Zhao et al. 2003). Ethidium is trapped in the nucleus by intercalating into DNA, leading to an increase of ethidium fluorescence. The cells (1 x 106/ml) were stained with dihydroethidium (1 µmol/l) by incubation in Tyrode’s buffer for 10 min, at room temperature, in the dark. After dihydroethidium labeling, cells were treated with the fatty acid mixture (0.1, 0.2, 0.3, and 0.4 mM) for additional 30 min. The same procedure was carried out in the presence of 0.2% BSA. Fluorescence was measured using FL-3 filter (excitation wavelength 480 nm/emission 670 nm) and analyzed in a FACS Calibur flow cytometer (Becton Dickinson, San Juan, CA, USA). Cells with dihydroethidium fluorescence were then evaluated using the Cell Quest software. Histograms of 10 000 events were analyzed per experiment.
NO measurement by DAF-2-FM fluorescence intensity assay
For this assay, cells were analyzed using a fluorescence microscope with 495/515 nm emission wavelength and KS 300 software (Zeiss, Munchen-Hallbergmoos, Germany). Lymphocytes were incubated in RPMI-1640 medium for 3 h in the presence or absence of the FFA mixture at 0.2 and 0.4 mM. Before fatty acid treatment, cells were loaded with DAF-2-FM (10 µM), pH 7.4, for 30 min, at 37 °C. NO-donor S-nitroso-N-acetyl-penicillamine (SNAP 6 nM), L-nitro-arginine (L-NA), and N (
)-nitro-L-arginine methyl ester (L-NAME 300 µM), as NOS inhibitors, or BSA (0.2%) were added to the medium as internal control. All experiments were performed at 37 °C in the dark to avoid non-specific DAF-2-FM oxidation.
Fluorescent measurement of intracellular calcium concentration ([Ca2+]i)
Changes in cytosolic calcium levels were fluorometrically monitored using the calcium-sensitive probe Fura 2-AM as previously described (Oliveira-Souza & De Mello-Aires 2000). The loading period for Fura 2-AM (15 µM) was 1 h in cells suspended in Tyrode’s solution. Afterwards, cells were treated with the fatty acid mixture at 0.2 or 0.4 mM and intracellular [Ca2+]i was monitored for 30 min. Fura 2-AM fluorescence was measured in 1 ml aliquots of cell suspension (1 x 107 cells/ml) using a Shimadzu model LS-5 fluorescence spectrophotometer set at 510 nm emission wavelength and excitation wavelengths alternating between 340 and 380 nm, with slit widths of 3 nm for excitation and 10 nm for emission. The cell suspensions were maintained at 37 °C and continuously stirred. A calibration procedure was performed at the end of each experiment. Transformation of the fluorescent signal to [Ca2+]i was performed by calibration with ionomycin (100 µM, maximum concentration) followed by EGTA addition (60 µM, minimum concentration) according to the Grynkiewicz equation, using the Kd of 224 nM (according to the Molecular Probes catalog). Under these conditions, mean control values of [Ca2+]i for lymphocytes from healthy subjects were 103.0 ± 8.3 nM (mean ± S.E.M. of six assays).
Proteolytic activities of caspases 3, 6, 8 and 9
Lymphocytes (1 x 106) were harvested after 24-h culture, centrifuged, washed in PBS, and subsequently lysed with 100 µl of the lysed buffer (100 mM HEPES–KOH, pH 7.5; 10% sucrose; 10 mM dithiotreitol (DTT); 0.1% CHAPS; 100 µM phenylmethylsulphonyl fluoride (PMSF); 10 µg/ml leupeptin; and 2 µg/ml aprotinin). Samples were incubated in ice for 30 min and centrifuged at 12 000 g for the same period. The soluble fraction was transferred to a new tube and protein was determined as described by Lowry (1976). Activities of caspases 3, 6, 8, and 9 were assayed following Thornberry (1998) with slight modifications. Briefly, 20 µg total protein were placed in 96-well plates. The reaction buffer (100 mM HEPES–KOH, pH 7.5; 10% sucrose; 10 mM DTT; 100 µM PMSF; 10 µg/ml leupeptin; and 2 µg/ml aprotinin) was then added at a final 100 µl volume. The plate was incubated at 37 °C for 15 min. Afterwards, 1 µl (50 µM) of the peptides Ac-DEVD-AMC, Ac-VEID-AFC, Ac-IETD-AFC, and Ac-LEHD-AMC as substrate for caspases 3, 6, 8, and 9 respectively, was added. The fluorescence of 7-amino-4-methylcoumarin and 7-amino-4-trifluoromethylcoumarin cleaved from Ac-DEVD-MCA, Ac-LEHD-AMC, Ac-VEID-AFC, and Ac-IETD-AFC, by caspases 3, 9, 6, and 8 respectively, was monitored for 30 min in a spectrofluorometer (Spectra Max Gemini XS, Molecular Devices, Union City, CA, USA) at 380 nm excitation and 460 nm emission.
Western blotting analysis
Lymphocytes (2 x 107) from control and fatty acid-treated cells were cultured for 24 h as described above. Afterwards, the cells were extracted and total proteins were separated by electrophoresis on SDS-PAGE followed by immunoblotting with anti-cytochrome c, anti-bid, anti-FasL, anti-14-3-3, anti-bad, and anti--actin primary antibodies. For investigation of cytochrome c release from mitochondrial inter-membrane space, lymphocytes were permeabilized by adding 100 µl extracting buffer (250 mM sucrose, 70 mM KCl, 137 mM NaCl, 4.3 mM Na2HPO4, 1.4 mM KH2PO4, 200 µg/ml digitonin, 100 µM PMSF, 10 µg/ml leupeptin, and 2 µg/ml aprotinin). Cells were then incubated for 30 min on ice and centrifuged at 1000 g for 5 min. The supernatant (100 µl) that contains cytochrome c was collected and 25 µl of 5 x SDS buffer (40 mM Tris–HCl, pH 6.8; 10% SDS; 50% glycerol; and 12.5% ß-mercaptoethanol) were added. The samples were then heated at 100 °C for 5 min and protein extracts were resolved on 15% SDS-PAGE.
For determination of FasL, 14-3-3, bid, and bad, total protein was extracted from the cells by adding 120 µl of previously heated extracting buffer (100 mM Tris-base, pH 7.5; 10 mM EDTA; 10% SDS; 100 mM sodium fluoride; 100 mM sodium pyrophosphate and 10 mM sodium ortovanadate). Cells were then sonicated for 1 min. Afterwards the samples were centrifuged at 12 000 g, at 4 °C, for 40 min. After centrifugation, the supernatant was utilized for protein determination. Just before the electrophoresis was carried out, the samples were heated at 100 °C for 5 min and protein extracts (30 µg) were resolved on 12% SDS-PAGE. After electrophoresis, proteins were transferred to nitrocellulose membranes (120 V, 1 h). The membranes were incubated with blocking solution (0.1% Tween 20, and 5% skimmed milk powder) for 2 h at room temperature followed by further incubation with renewed blocking solution containing cytochrome c (1:500), bid (1:500), FasL (1:350), 14-3-3 (1:350), and bad (1:350) for 4 h. The membranes were rinsed in washing solution (0.1% Tween 20, 0.15 mM NaCl, 20 mM Tris–HCl, pH 7.4; 3 x 10 min) and then incubated in a solution containing mouse anti-rabbit IgG horseradish peroxidase conjugated (1:10 000) for 1 h. Membranes containing immunorreactive bands were visualized by enhanced chemiluminescense (Amersham) according to the manufacturer’s protocol.
Protein determination
The protein content of the cell homogenates was determined by the method of Lowry (1976), using BSA as standard.
Statistical analysis
The results are expressed as mean and standard error of mean (S.E.M.) of 15 wells from at least three experiments, as indicated in the figures. ANOVA was employed to detect differences between the groups followed by the Tukey’s post-test. Student’s t-test was also employed to detect differences between the fatty acid treated and the control groups. P < 0.05 was taken to indicate significant differences.
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Fatty acid mixture at 0.2 and 0.4 mM increased DNA fragmentation, mitochondrial depolarization, and phosphatidylserine externalization in lymphocytes after 48-h culture as compared with the control group (Fig. 1
). The fatty acid mixture induced a dose-dependent increase of DNA fragmentation and phosphatidylserine externalization. Addition of BSA (0.2%) into the culture medium abolished this effect.
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Appropriate controls were carried out using fatty acid mixture at 0.1, 0.2, 0.3, and 0.4 mM in the assay plus lucigenin without cells. The fatty acid mixture used in the present study did not directly affect the lucigenin ROS-detecting system as previously described (Hatanaka et al. 2006). Kinetic studies showed that induction of superoxide production by human lymphocytes is a fast event that occurs within minutes after treatment of the cells with the fatty acid mixture (Fig. 2A). The use of fatty acid mixture raised the chemioluminescent signal indicating an increase in extracellular superoxide levels. Fatty acid mixture raised superoxide production by 3.5-, 9.9-, 17.3-, and 10.8-fold for 0.1, 0.2, 0.3, and 0.4 mM respectively, as compared with the control group. The increase observed was completely abolished by the addition of BSA to the culture medium (Fig. 2B). The inhibition caused by DPI, an NADPH oxidase inhibitor, indicates the participation of NADPH oxidase for superoxide anion production induced by fatty acids.
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The treatment of human lymphocytes with the fatty acid mixture increased the intracellular ROS levels as indicated by the reduction of dihydroethidium (Fig. 3). The fatty acid mixture caused a dose-dependent increase of the intracellular ROS content by 2.2-, 3.4-, and 4.9-fold respectively for 0.2, 0.3, and 0.4 mM fatty acid concentration as compared with control group. The increase observed in intracellular ROS levels was also blocked by the addition of 0.2% BSA to the culture medium in all fatty acid concentrations used.
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After a 3-h incubation period with the fatty acid mixture, cells were harvested, centrifuged, and resuspended in 25 µl PBS to be analyzed in a fluorescence microscope. Fatty acid mixture at 0.2 and 0.4 mM significantly increased NO production by human lymphocytes as indicated by DAF-2-FM fluorescence as compared with the control group (Fig. 4). The increase observed was higher at 0.4 mM (900 RLU) than at 0.2 mM (490 RLU) fatty acid. The treatment of cells with BSA (0.2%) significantly decreased the fluorescence induced by the fatty acid mixture. As expected, SNAP (positive control) and L-NA and L-NAME (negative controls) increased and decreased the DAF-2-FM fluorescence respectively.
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The effect of the fatty acid mixture on intracellular calcium levels was examined using Fura-2-AM. Treatment with 0.2 and 0.4 mM caused a robust intracellular calcium increase (~75-fold higher) as compared with the control group (Fig. 5). The increase in [Ca2+]i levels was sustained during 30 min of monitoring. BSA (0.2%) addition abolished the increase in intracellular [Ca2+]i levels induced by the fatty acids.
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The effect of fatty acid treatment on activities of caspases 3, 6, 8, and 9 in human lymphocytes after 24-h culture was examined. Lymphocytes treated with 0.4 mM fatty acid mixture presented 87% increase in caspase-3 activity when compared with the control group (Fig. 6). Caspase-6 activity was also significantly increased in lymphocytes by 0.4 mM fatty acid treatment (20%). Similar increase of caspase-9 activity was observed in lymphocytes treated with 0.2 mM (40%) and 0.4 mM (106%) fatty acid mixture, whereas there was no change in caspase-8 activity as compared with the control group.
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The expression of bid pro-apoptotic protein was significantly raised by the fatty acid treatment as shown in Fig. 7C. The increase was about 40% at 0.2 mM and 85% at 0.4 mM fatty acid concentration as compared with the control group. In the same manner, there was an increase of cytochrome c release to the cytosol as indicated by the bands obtained using the western blot assay. The bands of cytochrome c obtained in the mitochondrial fraction indicate a decrease of this protein in this organelle. The expression of 14-3-3, bad, and FasL proteins was not modified by the fatty acid treatment.
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Discussion |
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Mitochondria play an essential role in regulating cell death, which occurs when their membranes become permeabilized. Mitochondria possess an outer membrane in close communication with the cytosol and an inner membrane involved in energy transduction. Outer membrane permeabilization is regulated by bcl-2 family proteins, which control the release of proteins from the mitochondrial intermembrane space; these proteins then activate apoptosis (Yang et al. 1995). Inner membrane permeabilization is regulated by the MPTP, which is activated by calcium and oxidative stress and leads to bioenergetic failure (Armstrong 2006). Mitochondria are also a significant source of ROS, although the assumption that 1–2% of all O2 consumed ends up as ROS is likely to be overestimated (Brookes et al. 2004). The delicate balance between beneficial and harmful effects of free radicals is a very important aspect of living organisms and is achieved by mechanisms called ‘redox regulation’. The process of ‘redox regulation’ protects living organisms from various oxidative stresses and maintains ‘redox homeostasis’ by controlling the redox status in vivo (Droge 2002). Pro-apoptotic signals such as cytochrome c and Smac/Diablo are released from mitochondria as aresult of mitochondrial damage. Several mechanisms have been suggested as the cause of mitochondrial damage and release of pro-apoptotic signals including high levels of intracellular calcium that trigger the MPTP to open irreversibly (Maia et al. 2006).
Evidence is presented herein that induction of human lymphocyte apoptosis by the fatty acid mixture involves production of superoxide anion, and NO, increased intracellular [Ca2+]i, mitochondrial cytochrome c release, and activation of the caspase cascade. Apoptosis was observed by an increase in DNA fragmentation, mitochondrial depolarization, and phosphatidylserine externalization. These results are in agreement with our previous studies that showed a significant increase in apoptotic features after treatment of lymphocytes with the fatty acid mixture at 0.4 mM (Otton & Curi 2005). The apoptotic features examined in human lymphocytes were partially abolished by the treatment of cells with 0.2% BSA (Fig. 1
). There was no loss of membrane integrity in lymphocytes treated with the fatty acids as shown by propidium iodide (PI) assay and by acridine orange/ethidium bromide staining assay even after 48-h culture (data not shown).
The release of cytochrome c and other pro-apoptotic factors from mitochondria is a pivotal early event in the apoptotic cascade of many cell types (Antonsson 2004, Danial & Korsmeyer 2004, Green & Kroemer 2004, Reed et al. 2004). Once in the cytosol, cytochrome c and pro-caspase-9 bind the cytosolic protein apaf-1 and dATP to form apoptosomes that promote caspase activation (Liu et al. 1996, Wang 2001). The mechanisms by which pro-apoptotic factors are released from mitochondria early in apoptosis are not well understood. The MPTP in its original definition referred to an abrupt transition in permeability of the inner membrane that occurred when mitochondria were treated in vitro with calcium and reagents that increased oxidative stress. The phenomenon was suggested to be the result of the opening of the discrete protein pore (< 1500 kDa) in the mitochondrial inner membrane (Hunter & Haworth 1979). It has been speculated that MPTP in the inner membrane opens and causes swelling of the matrix space. As the inner membrane has a much greater surface area than the outer membrane, the ensuing swelling ruptures the outer membrane and spills cytochrome c and other pro-apoptotic proteins into the cytosol (Skulachev 1996, Brenner & Kroemer 2000, Kroemer & Reed 2000).
Despite strong evidence linking the MPT pore, cytochrome c release, and apoptosis, the precise mechanism of cytochrome c release is still unknown and is likely to be dependent on cell type, apoptotic stimulus, and precise cellular conditions. Scorrano et al.(2002) found that during bid-induced apoptosis, mitochondrial cristae underwent significant structural remodeling that was crucial for the release of cytochrome c. This remodeling was potently blocked with CsA (which potently inhibited MPTP opening) suggesting a link between the cristae remodeling and the MPTP. The MPTP can be considered an important signaling pathway leading to cytochrome c release, but its involvement in the physical mechanism of cytochrome c release is still under debate. Non-MPT pore-mediated mechanisms of cytochrome c release may exist (Lizard et al. 1995, Nunnari et al. 1997, Crompton 1999).
Cytochrome c release can occur in the absence of mitochondrial depolarization and without loss of outer membrane integrity. These findings suggest the existence of a more selective mechanism of permeabilization, e.g. formation of a pore in the outer membrane (Antonsson et al. 1997, Brenner & Kroemer 2000, Pavlov et al. 2001, De Giorgi et al. 2002, Chipuk et al. 2004, Green et al. 2004). This alternative mechanism is supported by the discovery of mitochondrial apoptosis-induced channel (MAC), a novel channel that forms in mitochondria early in apoptosis (Pavlov et al. 2001, Dejean et al. 2006). The notion that MAC provides the pathway through the outer membrane for release of cytochrome c is supported by data obtained through combining electrophysiology with microscopic, biochemical, and molecular approaches. Our results demonstrated that the fatty acid mixture induces mitochondrial depolarization that resulted in cytochrome c release and caspase-9 activation. This early apoptotic event in fatty acid-induced lymphocyte apoptosis activates the formation of apoptosome and then activation of caspases 3 and 6 and subsequently the apoptotic features observed in this study. Extrinsic pathway is probably not involved in fatty acid-induced lymphocyte apoptosis, since there was no significant difference in caspase-8 activity and FasL protein expression.
The involvement of calcium, ATP, and mitochondria for the process of apoptosis has been the subject of the present discussion. How can Ca2+, a physiological stimulus for ATP synthesis, become a pathological stimulus for ROS generation, cytochrome c release, and apoptosis? The primary role of mitochondrial Ca2+ is the stimulation of ox-phos that occurs at various levels, including allosteric activation of pyruvate dehydrogenase, isocitrate dehydrogenase, and -ketoglutarate dehydrogenase, as well as stimulation of the ATP synthase (complex V; Das & Harris 1990). Stimulation of the TCA cycle and ox-phos by Ca2+ would enhance ROS output by leading the whole mitochondrion towork faster and to consume more O2. In addition, Ca2+ stimulation of NO synthase (NOS) generates NO, which inhibits complex IV, and this would also enhance ROS production.
Our results showed that intracellular [Ca2+]i (Fig. 5) was significantly increased by treatment of lymphocytes with the fatty acid mixture. Intracellular calcium concentration increase could promote important modifications in lymphocyte mitochondria such as an increase in ROS levels, as showed by dihydroethidium and lucigenin assays (Figs 2 and 3), activation of NO synthase, and then increase in intracellular NO levels (Fig. 4). Arachidonic acid and other FFAs have been shown to elevate intracellular calcium levels (Packham et al. 1995, Draper et al. 2004). However, Maia et al.(2006) showed that apoptosis of C3HA cells induced by fatty acids did not involve calcium as a mediator of mitochondrial dysfunction.
In addition, Ca2+ stimulation of NO synthase (NOS) generates NO, which inhibits complex IV, and this enhances ROS generation. One important physiological role of NO is the regulation of mitochondrial ROS output. Thus, mitochondria can act as a ‘redox signaling box’, converting an NO production into a ROS signal. This signaling axis operates within a physiological window of NO concentrations, and disruption of this axis by pathological levels of NO is detrimental to mitochondrial ATP synthesis and cell function. Furthermore, NO in conjunction with high mitochondrial Ca2+ concentration can inhibit mitochondrial complex I and leads to ROS formation (Brookes et al. 2004).
Although this complex pathway that involves ROS, Ca2+, and NO are far away to be solved, it is clear up to now that fatty acids induce apoptosis of human lymphocytes by a mechanism that involves cytochrome c release from mitochondria, activation of the caspase cascade, and increase of bid protein content, superoxide and NO production, and of cytosolic calcium concentration. Which of these factors is the cause or consequence of apoptosis induced by fatty acid treatment remains to be elucidated.
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Acknowledgements |
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Received in final form 31 July 2007
Accepted 6 August 2007
Made available online as an Accepted Preprint 8 August 2007
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