Division of Maternal and Child Health Sciences, Ninewells Hospital and Medical School, University of Dundee, Dundee, Scotland, United Kingdom
Submitted 27 September 2004 ; accepted in final form 16 May 2005
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ABSTRACT |
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cancer; oxygen conformation; mitochondrial nitric oxide synthase; mitochondrial metabolism
Hypoxia evokes one of two conserved metabolic responses. The oxygen (oxy) regulation response conserves rates of ATP synthesis and demand close to aerobic levels through an activation of glycolysis (i.e., a Pasteur effect). Although this response sustains cell function independently of PO2, it is inherently limited by the supply of fermentable substrate (glucose) and the ability of the cell to tolerate the accumulation of toxic metabolic end products (lactate, H+). The oxy conformation response, on the other hand, minimizes these problems by suppressing ATP synthesis in coordination with reduced ATP consumption by vital cell processes. Thus hypoxia invokes a cessation of glycolysis (a so-called "reverse" Pasteur effect) that is accompanied by a reduction in ion transport activity, protein and RNA turnover, urea synthesis, and gluconeogenesis. The result is a state of metabolic arrest where endogenous metabolic substrate is conserved, metabolic end-product poisoning is minimized, transmembrane ion gradients are maintained, and both the proteome and high-energy phosphate pool are stabilized. Metabolic arrest is a key feature of naturally occurring dormant states such as estivation, hibernation, or freezing; however, several of the core physiological features of this response are also displayed in disease states whose etiology includes chronic hypoxia (20, 23, 24, 38, 44). The effectiveness with which cells survive these conditions depends on the degree to which metabolic demand is suppressed without an irreversible decline in integrity.
Recent evidence suggests that cancer cells display heightened sensitivity to fluctuation in PO2 and are capable of surviving prolonged and severe hypoxia by invoking a state of metabolic arrest (19, 20, 23). However, as yet, no mechanism has been identified which satisfactorily explains the oxy-conforming response. For almost 60 years, it has been recognized that nitric oxide (NO) rapidly and reversibly inhibits cytochrome c oxidase activity (55); however, the identification of a mitochondria-specific isoform of NO synthase (NOS) (17, 18, 49) has raised the possibility that mitochondrial NO synthesis may actively regulate oxidative phosphorylation in response to a wide range of physiological stimuli, including hypoxia. NO and O2 compete for binding to the binuclear center which is formed by the hemoprotein cytochrome a and probably the Cu
of cytochrome c oxidase (CCO) (53). Recent studies have determined that the steady-state NO production rate in submitochondrial particles increases from a range of 2040 to 80100 nM with a rise in [O2] from 20 to 200 µM (2). As the competitive inhibition of CCO by NO is determined by the moment-to-moment concentration of oxygen in the mitochondrial matrix, these values suggest that the O2-to-NO ratio is diminished by approximately two- to fourfold at 20 µM and is calculated to account for a 1625% inhibition of CCO activity in submitochondrial particles.
mtNOS has recently been identified as a posttranslationally myristoylated and COOH terminus-phosphorylated isoform of nNOS that is expressed in a wide range of tissues (14). As the KmO2 of this enzyme exceeds mean intracellular [O2] by approximately two- to fourfold [reported KmO2 values are 37, 40, and 73 µM O2 for the liver, kidney, and brain, respectively (2)], its activity would not be expected to rise without an exceptional increase in intracellular PO2 in vivo. However, because NO is
3 times more soluble than O2 in biological membranes and as the mitochondrion is a highly membranous organelle, it is conceivable that nonmitochondrial sources of this radical could determine electron transport chain activity independently from mtNOS. The aim of this study was therefore to determine whether the metabolic response of cancer cells to a gradual, physiological hypoxia (100123 mmHg; 12529 µM at 37°C) is governed by the production of NO. With the use of a lung adenocarcinoma cell line (A549), the results show that NO production rates rise during the transition to hypoxia and cause a suppression of mitochondrial membrane potential (
m). The corresponding cessation of ATP synthesis occurs in the absence of a Pasteur effect, perturbation in the adenylate phosphate pool, or overt cell death. These events are the hallmarks of a coordinated reduction of ATP turnover that is the cornerstone of hypoxic metabolic suppression.
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MATERIALS AND METHODS |
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Cell and spheroid culture. A549 and H441 lung carcinoma cell lines were purchased from the American Type Culture Collection and were used within 30 passages from purchase. Native, SV-40 transformed human bronchial epithelial (HBE) cells were a gift from Dr. A. Mehta (University of Dundee, Scotland) and were used within a similar number of passages. Primary cultures of rat alveolar type II (ATII) cells were prepared by elastase digestion and IgG purification as described elsewhere (31). All cell lines and primary cultured cells were maintained in Dulbecco's minimal essential medium (DMEM) supplemented with 10% fetal calf serum (FCS) and antibiotics. Fibronectin-coated culture dishes were used to promote ATII cell adhesion. For all experiments, culture PO2 was controlled by growing cells in a temperature-, gas-, and humidity-controlled MACS VA-500 microaerophilic workstation (Don Whitley Scientific, Shipley, UK) equipped with N2-flushable access ports and an airlock.
For spheroid culture, a nonadherent substratum was created by coating the wells of 24-well culture plates (Costar) with 200 µl of 0.5% agarose in serum-free DMEM. A549 cells (5 x 105) were seeded into each well and then incubated for 96 h at 37°C in the presence of 1 ml of DMEM-FCS. The distribution of hypoxia within the spheroid mass was determined by supplementing the medium with 200 µg/ml pimonidazole at ambient PO2. For experiments involving iNOS blockade, 100 µM L-NIL was added for the final 24 h of the 96-h incubation period. At the end of each experiment, spheroids (12 mm diameter) were washed once in PBS, embedded in Matrigel (GIBCO-BRL), and fixed for 2 h in PBS containing 10% formalin. The distribution of hypoxia and tyrosine nitrosylation was determined from 5-µm-thick paraffin-embedded sections using monoclonal antibodies to pimonidazole (1:100 dilution, Hypoxiprobe kit) and nitrotyrosine (1:50 dilution) and appropriate species-specific FITC secondary antibody conjugate (1:500 dilution). Single and composite FITC and DAPI-stained images were obtained using a Zeiss Axioskop fluorescent microscope equipped with a Hamamatsu C4742-95 color digital camera using Openlab (Improvision, Coventry, UK) software. The number of intact nuclei and oligonucleosomes per mm2 was calculated by counting individual nuclei and fragmented chromatin in DAPI-stained sections by laying a grid composed of 40-µm2 squares over cross-sectional images of each spheroid.
For all experiments, care was taken to ensure that the media were preequilibrated to the appropriate oxygen tension and temperature.
Phosphorothioate-antisense ODN treatment. The protocol for antisense treatment of monolayers was as described elsewhere (22). Cells (1.5 x 105/well) were placed into monolayer culture in 6-well culture plates. After 24 h, the medium was replaced with 1 ml/well of DMEM containing 0.5% FCS and antibiotics as before. After a further 4 h, 100 µl of a transfection mixture were added that contained 20 µg/ml Lipofectamine 2000 and 1 µM of either random sequence control (±FITC tag) or iNOS AS-ODN that had been previously combined according to the manufacturer's instructions. Cells were left to incorporate antisense for 24 h before the start of each experiment. Knockdown of iNOS protein expression was confirmed by Western blot and immunofluorescence analysis, as detailed in RESULTS.
Preparation of mitochondria. Mitochondria were prepared from pooled A549 monolayer cultures using a Mitochondrial Fractionation Kit (Active Motif, Rixensart, Belgium) coupled with crude purification by differential centrifugation. The mitochondrial pellet was purified further by centrifugation through a discontinuous Percoll gradient (24/40%) at 14,000 g for 10 min at 3°C. Purified mitochondria were washed free of Percoll and placed in mitochondrial suspension buffer containing (in mM) 400 sucrose, 70 Na+-HEPES (pH 7.6), 100 KCl, 3 EDTA, 6 EGTA, 1 mg/ml aprotinin, and 1% BSA.
Fluorescence detection of mitochondrial NO synthesis and m.
Production of NO was followed using the cell-permeant NO-excited fluorescent dye DAF-FM diacetate. For studies conducted in isolated mitochondria, 10 mg (wet wt) of mitochondria were loaded with 10 µM DAF-FM diacetate for 1 h in the dark at ambient PO2 and 37°C. After being washed, rotenone (1 µM) and myxothiazol (1 µM) were added to inhibit endogenous reactive oxygen species (ROS) produced during the transition to hypoxia. Mitochondria were placed into a 5-ml quartz cuvette containing a stir bar, which was then mounted into the temperature-controlled recording cell of a Hitachi F-2500 fluorescence spectrophotometer. Acute deoxygenation was achieved by flushing the interior of the recording chamber with a hydrated gas mixture containing 3% O2-5% CO2-balance N2 and by the equilibration of PO2 in the cuvette to the gas mixture was measured simultaneously using an ISO2 oxygen meter. Preliminary experiments indicated that this method produced a steady rate of deoxygenation in the cuvette without artefactual change in the production of DAF-FM fluorescence. In each case, NO production observed as DAF-FM fluorescence was monitored during deoxygenation using an excitation wavelength of 495 nm and after light was emitted at 515 nm.
m was measured in intact cells using the J-aggregate-forming lipophilic cation JC-1. Cells were incubated in the dark at 37°C in the presence of 3 µM JC-1 at PO2s of either 23 or 100 mmHg using the microaerophilic workstation. After incorporation, cells were washed free of excess dye with O2-equilibrated serum-free DMEM. The cells (25 mg wet wt) were placed into a 5-ml quartz cuvette containing a stir bar and sealed using a gas tight rubber stopper under an atmosphere containing the correct PO2.
m was measured by following the change in emitted fluorescence at 525 and 590 nm on excitation at 488 nm. Additions of the NO donor 2,2'-[hydroxynitrosohydrazono]bis-ethanimine (DETA-NO) or the NOS inhibitor L-NAME were made through an injection port in the top of the gas tight cap.
For studies examining the effects of hypoxia on NO release and m in cancer-derived (A549, H441) and native tissue-derived cells (HBE, rat ATII) (Fig. 10), cells were grown to confluence in 24-well culture dishes at ambient PO2. Simultaneous measurements of NO release and
m during the transition from room air to anoxia were made in DAF-FM- and JC-1-loaded cells in serum-free Hanks' balanced salt solution containing (in mM) 20 Na+-HEPES (pH 7.4), 137 NaCl, 5.36 KCl, 1.26 CaCl2, 0.49 MgCl2, 0.41 MgSO4, 0.44 KH2PO4, and 0.34 Na2HPO4 and 5 glucose, using a BMG-Labtech Fluostar Optima fluorescent microplate reader equipped with a gas port, microinjector, and temperature control. To initiate the transition toward hypoxia, the measuring chamber of the reader was flushed with a gas mix containing 95% N2-5% CO2. Dissolved PO2 in each well was measured by following the change in fluorescence emission at 525 nm of the O2 quenched dye dichloritris(1,10-phenantholine)-ruthenium (II) hydrate [Ru(phen)32+, 100 µM] (12). Dye calibration was achieved within the plate reader at the start and finish of each experiment by observing the fluorescence obtained in media equilibrated to 0 and 142 mmHg. Measurements were taken at 60-s intervals throughout the transition to anoxia. Where described, gas-equilibrated iodoacetate was added to each well at the given concentration during the experimental run via the microinjector. Cell viability was determined at the end of each experiment by addition of 0.04% (vol/vol) Trypan blue for 10 min, followed by three washes in PBS. Staining intensity was determined in each well by measuring light absorbance at 595 nm with background correction at 492 nm. Protein concentration in each well was determined using the Bio-Rad Protein Assay.
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m was examined in spheroids by the same approach. Spheroids were grown to
100-µm diameter and were then incubated in suspension with 10 µM JC-1 for 2 h, after which they were washed three times in serum-free DMEM. Spheroid suspension (200 µl) was then placed into each chamber on a Labtek (Nalge, Naperville, IL) chambered coverslip and 150-µm Z-stack images were obtained in the presence or absence of 100 µM L-NIL under identical laser power and gain settings. Qualitative comparisons were made by comparing the distribution of the J-monomer and aggregate in optical slices obtained 30 µm in from the base of the spheroid. Homogeneous distribution of the J-monomer confirmed even uptake of the dye throughout the miniature spheroid.
Detection of tyrosine nitrosylation, NOS isoforms and activity. Nitrosylation of tyrosine residues was determined by Western blot analysis of mitochondrial proteins using an anti-rabbit nitrotyrosine antibody (1:2,000 dilution). Parallel blots were treated with 100 mM Na2S2O4 overnight at 4°C to convert nitrosylated residues to their antibody-blind amino form. The expression and cellular location of NOS isoforms were determined by Western blot analysis and immunofluorescence. Western blotting was performed as described before (39). Neuronal NOS (nNOS), iNOS, and endothelial NOS (eNOS) were identified using anti-rabbit NOS 1-R20 (sc-648), NOS 2-M19 (sc-650), or NOS 3-C20 (sc-654) each at 1:1,000 dilution with secondary detection by a species-specific HRP antibody conjugate. Positive controls were LPS-treated RAW 264.7 macrophage protein for iNOS and a mouse brain homogenate for nNOS and eNOS. For immunofluorescence, cells were grown in Lab-Tek chambered coverslips and maintained in normal monolayer culture conditions at PO2s of either 100 or 23 mmHg or removed from 100 to 23 mmHg for 1 h. Cells were washed in PBS, fixed for 10 min in 10% neutral PBS-buffered formalin, and then permeabilized by incubation for a further 15 min in PBS containing 0.1% Triton X-100. Nonspecific binding was minimized by being blocked in 5% rabbit serum for 1 h followed by incubation with primary antibodies to iNOS, nNOS, or eNOS each at 1:50 dilution overnight at 4°C. Secondary detection was performed using anti-rabbit or anti-goat IgG-FITC (NOS isoforms) or anti-mouse TRITC (cytochrome c) at 1:500 dilution. Chambered coverslips were examined by laser confocal microscopy (model LSM-510; Zeiss, Gottingen, Germany), and images were acquired and analyzed using Zeiss Image Examiner software.
RT-PCR was used to determine the distribution of iNOS mRNA in cancer and native tissue-derived cell cultures. Reverse transcription was performed on 1 µg of RNA with the subsequent PCR reaction performed on 10 ng of cDNA. Reactions were also routinely performed without reverse transcription to control for possible contamination of RNA by genomic DNA. The primers used spanned the proximal 3' untranslated region of human iNOS mRNA (GenBank accession no. NM-153292 and were sense (nt 36163635), ACAGGAGGGGTTAAAGCTGC, and antisense (nt 38293847), TTGTCTCCAAGGGACCAGG, yielding a product size of 231 bp. For rat iNOS (Genbank Accession No. U26686), the primers used were sense (nt 12321251), CTGTCACCGAGATCAATGCA, and antisense (nt 18901909), CATGAGCAAAGGCACAGAAC, yielding a 677-bp product. The identity of bands resolved at the correct molecular weight was confirmed by sequencing.
NOS activity was determined using 5 µg of mitochondrial protein placed into a buffer containing (in mM) 66.7 Tris (pH 7.4), 2.7 NADPH, 3.3 CaCl2, 1 ornithine, 0.01 BH4, 0.002 FMN, and 0.002 FAD at 37°C. Where necessary, the buffer had been previously equilibrated to PO2s of 100, 40, 23, or 7 mmHg, and Ca2+-free conditions were achieved by omission of CaCl2 from the reaction mixture together with the addition of EDTA (5 mM). Reactions were initiated by addition of 0.5 µCi L-[U-14C]arginine and allowed to proceed for 30 min at 37°C at the respective PO2 using the microaerophilic workstation. Reactions were terminated by addition of 1 ml of ice-cold stop buffer containing (in mM) 30 Na+-HEPES (pH 5.5) and 3 EDTA. NOS-generated radiolabeled citrulline was separated from arginine by passage over an ion-exchange column consisting of 0.5 ml of Dowex-50 W resin (Na+ 50X8-400, Sigma). Citrulline was eluted by addition of a further 1 ml of stop buffer, and radioactivity was detected by the addition of 20 ml of Optiphase "Trisafe" scintillant (Perkin Elmer, Loughborough, UK) followed by detection using a liquid scintillation counter (Canberra-Packard, Pangbourne, UK). Background counts were measured in parallel in the presence of 1 mM L-NAME and were subtracted from the total count yield from each assay.
Measurement of CCO activity in gas-equilibrated whole cell homogenates.
All manipulations were conducted using buffers equilibrated to the appropriate PO2 using the MACS VA 500 microaerophilic workstation. Cells grown to 90% confluence at PO2s of either 23 or 100 mmHg were placed into the appropriate gas-equilibrated assay buffer composed of (in mM) 250 sucrose, 120 KCl, 10 Tris·HCl (pH 7.0), and homogenized (3 x 30 s) on ice using an Ultra Tarrax homogenizer at full speed. Enzyme activity in each homogenate was assessed using the Cytochrome c Oxidase Assay Kit (Sigma) by following the increase in Na+ hydrosulfite-reduced ferrocytochrome c substrate absorbance at 550 nm over 60 s. The PO2 of each assay reaction was conserved during each reaction by sealing the cuvette with a gas tight rubber stopper inside the MACS VA500 workstation with additions of gas-equilibrated DETA-NO or L-NAME made through an air-tight port. CCO activity was calculated assuming mM (ferro-
ferricytochrome c) = 21.84.
Measurement of metabolites, metabolic activity, apoptosis, and necrosis. Rates of glucose oxidation and fermentation to lactate were measured in A549 cells grown to confluence in 25-cm2 culture flasks. Before the experiment, the medium was exchanged for gas-equilibrated Hanks' balanced salt solution + 5 mM glucose, and the cultures were placed at the appropriate humidified gas atmosphere. Glucose oxidation rates were measured by the addition of 0.1 µCi D-[U-14C] glucose (303 mCi/mmol) to the culture medium for 2 h and sealing the flasks with an airtight stopper. At the end of the incubation, 0.5 ml of 7% HClO4 was injected through a gas tight seal in the top of each flask to liberate 14CO2 into the atmosphere and total CO2 was collected by pumping four volumes of the flask atmosphere through 2 ml of hyamine hydroxide solution. The concentration of 14CO2 was assessed by performing liquid scintillation counting using a Canberra-Packard scintillation counter. Glucose fermentation to lactate was measured by following the rate of 3H2O liberation from D-[5-3H(N)]-glucose. A549 cells were treated identically to the glucose oxidation experiments with the exception that 0.1 µCi D-[5-3H(N)]-glucose (20 Ci/mmol) was added to each flask at the start of the experiment. After 2 h, the medium was removed and 3H2O generated during glycolysis was separated from D-[5-3H(N)]-glucose by applying 1 ml of medium to a 5 x 1-cm-diameter column containing Dowex 1-X4 anion exchange resin (200400 mesh) preequilibrated in H2O. Using a [14C]glucose tracer placed into the experimental medium, we determined that the columns retained 94 ± 0.7% of the radioactivity associated with glucose on the column.
[ATP], [ADP], and [AMP] were measured in neutralized perchloric acid cell extracts using a Knauer K-1001 high-pressure liquid chromatography pump fitted with a 15 x 4.6-cm Supelcosil LC-18-T column (3 µm particle size) (Supelco, Bellefonte, PA). Adenylate phosphates were independently resolved by perfusing the column with either buffer A, composed of (in mM) 100 KH2PO4, 4 tetrabutylammonium hydrogen sulfate (pH 6.0), or buffer B, composed of 70% buffer A plus 30% methanol (pH 7.2) according to the following gradient protocol operated at a flow rate of 1 ml/min [time (min), %buffer B]: 0, 0; 2.5, 0; 5, 30; 10, 60; 13, 100; 18, 100; 19, 0. Detection of metabolites eluted from the column was achieved using a Knauer 2500 UV detector. Peak areas were converted into concentration by application of the data to standard curves determined over a range of 0100 nM for each metabolite.
Lactic acid and glucose were measured according to the method of Bergmeyer (6), and nitrite production was determined using the Greiss assay. In each case, results were normalized to cellular protein content. The incidence of apoptosis and necrosis were determined by oligonucleosome and lactate dehydrogenase release, as described before (39).
Statistics. Data were analyzed using one-way ANOVA with post hoc significance assessed with Tukey's honestly significant difference test using statistical software (SigmaStat Jandel, version 3.02, San Rafael, CA). Values are given as means ± SE. A P value <0.05 was considered to be statistically significant.
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RESULTS |
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When subjected to a gradual change from ambient PO2 to 23 mmHg, isolated mitochondria showed a hypoxia-dependent increase in DAF-FM fluorescence that was abolished by L-NAME (Fig. 1A). The pooled rates of DAF-FM fluorescence expressed as a proportion of the endogenous rate of change at ambient PO2 (Fig. 1B) showed the hypoxic rate of NO production at 23 mmHg was fourfold greater than the ambient rate but remained raised at PO2s approaching anoxia (<1 mmHg). Ca2+-independent NOS activity in submitochondrial particles, determined as the L-NAME-inhibitable rate, increased over the same PO2 range as the rise in DAF-FM fluorescence, reaching a maximum from 4823 mmHg (Fig. 2A). The rate returned to prehypoxic levels at PO2 = 7 mmHg. Further investigation in isolated mitochondria revealed evidence for both Ca2+-dependent and -independent isoforms (Fig. 2B). In normoxia, the rate in the presence of Ca2+ was threefold greater than the equivalent hypoxic rate but was neither enhanced by calmodulin nor abolished by removal of Ca2+ or L-NIL. In hypoxia, calmodulin raised the baseline Ca2+ rate by 3.5-fold to match the activity found in normoxia. However, the largest increase in activity occurred in hypoxia under Ca2+-free conditions (Ca2+-free buffer + 5 mM EDTA) where the hypoxia activated rate was threefold greater than the corresponding normoxic activity and was entirely abolished by the iNOS inhibitor L-NIL.
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Western blots of whole cell lysates showed nNOS and iNOS to be present in A549, but there was no immunoreactivity for eNOS (Fig. 3A). Confocal immunofluorescent images (Fig. 3B) revealed a punctate distribution of both the anti-nNOS and anti-iNOS-FITC signal, which strongly colocalized with anti-cytochrome c-TRITC. A similar distribution for both NOS isoforms was observed in H441 and HBE cells, and in each case, culture PO2 had no observable effect on the distribution or staining intensity of either NOS isoform (data not shown).
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NO production, hypoxia, and cell survival in tumor spheroids. A549 tumor spheroids were used to determine whether hypoxia was linked to NO production and cell survival in a densely packed cell population reminiscent of the solid tumor core. Pimonidazole, a hypoxia marker, was consistently confined to the core (C) region of the spheroid (Fig. 11A), indicating that a hypoxic gradient extended from the periphery (P) to the center of the spheroid. This was associated with an increase in the number of oligonucleosomes and intensity of tyrosine nitrosylation within the core region. The addition of 100 µM L-NIL to tumor spheroids resulted in the fragmentation of chromatin within the core region, loss of structure, and a reduction in nitrotyrosine-positive antibody staining. Figure 11B shows a change in the total number of intact nuclei vs. oligonucleosomes under control conditions or in the presence of L-NIL. In each case, the oligonucleosome fraction in the core region increased significantly by 2.5- and 5.5-fold, respectively. Figure 11C shows the distribution of J-monomer and aggregate in JC-1-loaded spheroids cultured at ambient PO2. Although the distribution of the J-monomer (green) indicated that the dye was evenly distributed within the spheroid, J-aggregate fluorescence (red), indicative of actively respiring mitochondria, was confined to the oxygenated spheroid periphery. The addition of 100 µM L-NIL altered the pattern of J-aggregate fluorescence such that overall fluorescence intensity was increased and there was a wider distribution of fluorescence within the spheroid core. These results are typical of five independent observations.
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DISCUSSION |
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Taking each point in turn, mitochondrial production of NO in A549 cells was acutely oxygen sensitive, rising to a rate that was 4-fold greater in hypoxia compared with normoxic controls. This was matched by an increase in L-NAME-inhibitable NOS activity that was maximal between PO2s of 40 and 23 mmHg and which showed a mixed catalytic dependency for Ca2+. In normoxia, the rate of NOS activity in the presence of Ca2+ was threefold greater than the hypoxic rate but was not increased further by calmodulin and showed no statistically significant change regarding the removal of Ca2+ or the addition of the iNOS inhibitor, L-NIL. In hypoxia, however, both Ca2+-dependent and -independent activities were present as calmodulin activated the rate in the presence of Ca2+ alone by 3.5-fold but complete removal of Ca2+ raised NOS activity to a level that was 6-fold greater than the Ca2+ rate. Significantly, this effect was almost entirely abolished by the iNOS inhibitor L-NIL. These data suggest that two NOS isoforms are present within mitochondria of A549 cells: one Ca2+ dependent and inhibited by hypoxia (Fig. 2 and Ref. 2) and the other Ca2+ independent and induced by hypoxia.
Although the identity of mitochondrial NOS remains controversial (e.g., 37), recent studies conducted in mouse NOS-knockout models (32) have provided support for the original observation by Elfering et al. (14) that this enzyme is a Ca2+-activated splice variant of nNOS. In the present study, immunoreactivity for both nNOS and iNOS strongly colocalized with cytochrome c, indicating that both isoforms are present in mitochondria. In accord with the NO synthesis data from isolated mitochondria that suggests a Ca2+-independent, L-NIL-inhibitable NO source is active in hypoxia, confocal studies of intact cells showed increases in both cytosolic and compartmented DAF-FM fluorescence occurred during the hypoxic transition that were diminished by iNOS antisense ODN. Given the target specificity of antisense ODN, this observation identifies the iNOS isoform as the agent of hypoxic NO production in A549 cells.
iNOS has been reported to localize with other NOS isoforms in cardiac muscle mitochondria (56) and occurs as an inducible component of mtNOS activity in endotoxin-exposed lung (9, 10, 15). Our report of an increase in mitochondria-associated iNOS activity during the early-to-midddle phases of the hypoxic transition accords with these observations and provides a mechanism that could underpin the tendency of cancer cells to oxy-conform. However, because oxygen is a compulsory substrate for the conversion of arginine to citrulline, this mechanism supposes that falling oxygen availability does not limit iNOS activity during either the hypoxic transition or the steady state. Reports on the KmO2 of iNOS show little consensus. Values range from 5 mmHg for the isolated enzyme to 65 mmHg in both activated macrophages and the intact lung (13, 43, 47). Other studies show O2 substrate limitation is apparent only between PO2s from 7.9 to 39.4 mmHg (apparent KmO2 = 14 mmHg) (45) and that iNOS-dependent signaling to soluble guanylate cyclase is not altered by severe hypoxia (32 mmHg) (3). Our own study showed that NOS enzyme activity was induced between PO2s of 4023 mmHg and was diminished at 7 mmHg (Fig. 2A), suggesting that relatively severe hypoxia (<23 mmHg) is necessary to limit iNOS function. Nevertheless, NO production rate (as DAF-FM fluorescence) was significantly greater than the aerobic rate at PO2s of <1 mmHg (Figs. 1 and 10B) and remained so despite diminished iNOS activity. A similar disparity between NOS activity and NO production has been reported to occur in activated macrophages at PO2 <7.9 mmHg (45) and has been suggested to arise from anaerobic, NOS-independent pathways, which reduce nitrite to NO. Indeed, we observed a positive relationship between depth of hypoxia and iNOS-dependent nitrite accumulation in solution (Fig. 8B), which is consistent with the generation of a substrate source for these pathways. Recent work (40) has identified xanthine oxidase as an effective reducer of organic/inorganic nitrate/nitrite that is capable of producing NO and sustaining soluble guanylate cyclase activity in anaerobic conditions. Anaerobic nitrite reductase activity has also been shown to occur in mitochondria through respiratory chain-dependent (nitrite cycling at the ubiquinol oxidant site of cytochrome bc1 complex III; Ref. 36) and independent (aldehyde dehydrogenase 2 activity; Ref. 34) pathways. In addition to these potential sources of anaerobic NO production, hypoxia potentiates the synthesis and biological effects of NO by increasing iNOS genomic expression and raising the bioactive half-life of NO through the cessation of NO consumption and oxidation (27, 42, 52). These observations therefore support a model in which NOS-dependent and -independent NO
NO2 cycling, coupled with the tendency of hypoxia to increase NO bioactivity, sustains the physiological effects of NO at any depth of hypoxia, including anoxia.
A central aspect of this model concerns the mechanism of iNOS activation during the middle phases of the hypoxic transition. iNOS activation involves the relief of inhibitory protein [kalirin, NOS-associated protein 110 kDa (NAP110)] interactions with the iNOS monomer, disulfide bridge formation between iNOS monomers and the subsequent binding of zinc and biopterin to form a stable, catalytically competent homodimer. Further potentiation of enzyme activity involves Rac1 and Rac2 GTPase-dependent intracellular trafficking and, particularly in epithelial cells, confinement of iNOS activity at specific membrane locations by interaction with the ezrin-radixin-moesin-binding phosphoprotein 50 (48; reviewed in Ref. 57). Given the potential for intracellular redox state to modulate disulfide bridge formation, it is possible that altered ROS production during the hypoxic transition could modify this process. Although mitochondrial ROS production increases transiently in A549 cells during the transition to hypoxia (38), blockade of mitochondrial ROS synthesis with rotenone and myxothiazol did not affect the hypoxic rise in NO production (Fig. 1). We also examined the possibility that translocation toward the plasma or mitochondrial membranes could potentiate the activity of iNOS in hypoxia by exposing the enzyme to a region of high O2 solubility. However, we found no effect of PO2 on the proportion of iNOS that colocalized with cytochrome c (data not shown). Similarly, there was no significant redistribution of the enzyme toward the apical membrane as has been found to occur in intestinal epithelial cells exposed to flagellin (48). In addition, an iNOS-like enzyme activity could be induced by hypoxia in mitochondria that had been isolated from normoxic cells (Figs. 13). Although the activation mechanism remains unclear, these data suggest that the hypoxic increase in iNOS activity was endogenous to mitochondria, did not require mitochondrial ROS production and did not involve overt translocation of cytosolic iNOS toward the mitochondrial membrane. We also note that the lack of inhibition of mitochondrial NO production by rotenone and myxothiazol in severe hypoxia (Fig. 1) eliminates respiratory chain nitrite reductase activity as a potential NOS-independent source of NO under these conditions (36).
The third line of evidence in support of a role for NO as a mediator of hypoxic metabolic arrest stems from the effects of NO production on m. Beltran et al. (4) reported that blockade of complex IV by exogenous NO results in a time-limited conservation of
m in lymphoid T cells that is sustained through an increase in glycolytic ATP production (a Pasteur effect) and reversal of the F1F0 ATPase. They argued in favor of this mechanism as a means of protecting the cell against apoptosis and suggested that it underpins the anti-apoptotic effects of NO. These data are supported by further studies in astrocytes and neurons that showed that glycolytic activation and conservation of
m during NO inhibition of complex IV is cell specific and requires activation of AMP protein kinase by NO and subsequent phosphorylation of 6-phosphofructo-2 kinase (1). However, our results in A549 cells showed that the partial dissipation of
m in hypoxia was NO dependent and was not accompanied by either a Pasteur effect or a change in high-energy phosphate concentrations (Figs. 7 and 8). Moreover, direct measurements of apoptosis and necrosis did not indicate any statistically significant change in cell viability with time spent in hypoxia (Fig. 9). Blockade of NO synthesis by L-NAME and iNOS antisense ODN profoundly reversed these effects by raising glucose consumption and lactate production severalfold, maintaining
m at normoxic levels, increasing normoxic ATP concentrations, destabilizing the high-energy phosphate pool in hypoxia and causing a significant increase in cell necrosis over 24 h. A similar requirement for NO production in maintaining cell viability in hypoxia was found in cancer-derived H441 cells and native tissue-derived transformed HBE cells, both of which expressed iNOS and displayed elevated rates of NO production in hypoxia (Fig. 10). In ATII cells, where iNOS expression and NO production were absent,
m in hypoxia was intrinsically dependent on glycolytic ATP production as shown by a lethal depolarization (indicated by a failure to exclude Trypan blue), which occurred in the presence of IAA. Taken together, these results suggest that NO produced during the hypoxic transition limits the scope of hypoxic glycolytic activation (i.e., the Pasteur effect), stabilizes
m to a less-energized state and suppresses the rate of cellular ATP utilization during hypoxia in cancer cells.
In the absence of a Pasteur effect, the conservation of both high-energy phosphate concentrations and cell viability can prevail in the face of reduced mitochondrial ATP synthesis only by a coordinated reduction in ATP synthesis and demand (i.e., metabolic arrest); therefore, these observations support an alternative cytoprotective role for NO that is in contrast to the glycolytic ATP synthesis rescue model proposed by Beltran et al. (4). The differences between the two strategies may well reflect different environmental characteristics of the cell systems in which these phenomena have been investigated. Thus, in astrocytes, neurons, and lymphoid T cells, which are likely to experience hypoxia and nutrient restriction only in pathophysiological states, the compensatory glycolysis evoked by NO during complex IV blockade may be sufficient to sustain m for a period of several hours. However, the efficiency of this process over time is inevitably limited by the availability of fermentable substrate (glucose) and is open to self-poisoning by the accumulation of toxic metabolic end products (lactate and H+). In cancerous cells, however, the severely hypoxic and nutrient-deprived environment within the solid tumor core creates a strong selective pressure favoring those cells that are adapted for surviving long periods of oxygen or glucose restriction. Metabolic arrest represents by far the most secure strategy for survival under these conditions because fermentable substrate supplies are conserved, the production of metabolic end products is limited, and the intracellular milieu is inherently stabilized by arrest of protein and RNA turnover, so the period of survival becomes less dependent on the duration of the stress.
To determine whether this form of metabolic regulation could, in principle, operate in the solid tumor core, A549 tumor spheroids were used to recreate the oxygen- and nutrient-restricted conditions, which typifies this environment (Fig. 11). An extensive hypoxic core region was identified by pimonizazole staining, which shared a common distribution with the binding of antibodies against nitrotyrosine, a marker of NO production. In microspheroids, the hypoxic core region was also found to display suppressed mitochondrial activity, which was raised throughout the spheroid mass during iNOS inhibition. As with monolayer cultures, where iNOS inhibition evoked a dramatic rise in necrotic lysis within 24 h (Fig. 9), cells within the spheroid core showed widespread evidence of nuclear fragmentation following the same treatment, suggesting that in this hypoxic environment, loss of NO production triggers cell clearance by apoptosis. Thus NO production in the spheroid core is associated with suppressed mitochondrial function and critically maintains cell viability.
Although the spheroid model we used here is simplistic, it accords with other studies that report raised NOS activity and NO production in several types of tumors (reviewed in Ref. 7). The role of NO in tumorigenesis has been difficult to establish, however, as its actions are variously cytoprotective/proliferative or cytotoxic, depending on the site, production rate, concentration, and metabolic fate of NO (7, 30, 51, 52). Thus, at low concentrations, NO is anti-apoptotic, proliferative, pro-angiogenic, and contributes toward mutations in cell cycle-regulating genes (e.g., p53), but at higher concentrations it induces position-dependent cell cycle cytostasis, p53 accumulation, and DNA damage, including p53 mutation and apoptosis (30). Although our studies did not establish the molar concentration of NO required to initiate metabolic arrest, the appearance of nitrotyrosine residues among mitochondrial proteins and within the hypoxic spheroid core is consistent with relatively high NO production rates necessary for cytostasis. During hypoxia, NO-evoked metabolic arrest could play a crucial role in maintaining cell integrity during cytostasis by suppressing pro-apoptotic signals. Such a mechanism would effectively conserve a population of viable cells in the tumor core, which would be capable of metastasis on the reintroduction of oxygen and nutrient-rich conditions.
This study implicates NO as an important signaling agent during oxy-conformation in cancer cells and accords with several lines of evidence that suggest that NO profoundly influences cellular adaptation to hypoxia. One consequence of the competitive binding of NO to the binuclear catalytic center of CCO is to reduce the mitochondrial O2 diffusion gradient toward mitochondria and thus redistribute O2 toward nonrespiratory oxidases. This effect accounts for the tendency for low NO concentrations (<400 nM) to destabilize the tumorigenic transcription factor, hypoxia-inducible factor-1, in hypoxia by activation of O2-dependent prolyl hydroxylases (21), whereas higher concentrations have the opposite effect (41, 51). Hence, at permissive concentrations, NO decentralizes respiratory control from the mitochondrion to the cytosol and thus profoundly alters the response of oxygen-sensing pathways to changing PO2. Aside from NOs effects on CCO, NO-dependent S-nitrosylation and ADP ribosylation have also been shown to inhibit electron flow through other respiratory complexes [e.g., complex I (5)] and block the activity of rate-limiting glycolytic enzymes [e.g., glyceraldehyde-3-phosphate dehydrogenase (5, 35)]. Hypoxia and NO induce tolerance to glucose starvation in normal and cancerous cell lines by activation of 5'-AMP-activated protein kinase and protein kinase B/Akt (16). However, as discussed above, these studies are offset by others, which show that NO activates glycolysis during respiratory arrest in cell lines that are vulnerable to hypoxia (1). More generally, tyrosine nitrosylation, found here to be induced in mitochondrial proteins during hypoxia and in the spheroid core, may fundamentally alter protein structure with consequences for enzyme catalytic behavior (reviewed in Ref. 28).
Perhaps the most widely described effect of NO, which is consistent with metabolic suppression, concerns the inhibition of anabolic processes. In Drosophila, Teodoro and O'Farrell (50) showed that NO precisely mimics the effects of hypoxia by causing an inhibition gene expression and protein turnover. Their results identify NO as a crucial signaling agent that causes the reversible arrest of Drosophila development during hypoxia. In cancer, NO is widely reported to induce a long-lasting cytostasis in the G1 phase that involves a rapid and reversible inhibition of ribonucleotide reductase with downregulation of cyclin D1 expression, a crucial component for cell cycle progression (46). In parallel with these effects, NO inhibits protein synthesis at the level of translation initiation, possibly through the activation of a eukaryotic initiation factor-2 kinase or inhibition of a phosphatase (11, 33). In coordination with a reduction in mitochondrial function and glycolysis, the suppression of DNA and protein synthesis by NO is likely to constitute a major element of the metabolic suppression reported in this study.
In conclusion, this study has shown that hypoxia evokes an iNOS-dependent production of NO in mitochondria that causes a reduction in mitochondrial ATP synthesis without compensation by a Pasteur effect and that is coordinated with a suppression in ATP demand. As a consequence of this hypoxic metabolic suppression, the survival characteristics of cells within the solid tumor microenvironment are extended and likely contribute to the malignancy of these tumors on reoxygenation. These results highlight NO as a key regulator of this phenomenon and suggest that targeting of iNOS expression in solid tumors may render them amenable to conventional chemo- and radiotherapy techniques.
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ACKNOWLEDGMENTS |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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