Department of Medicine, Indiana University and Roudebush Veterans Affairs Medical Center, Indianapolis, Indiana 46202
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ABSTRACT |
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Guanine nucleosides are toxic to some forms of cancer. This toxicity is pronounced in cancers with upregulated guanine nucleotide synthesis, but the mechanisms are poorly understood. We investigated this toxicity by measuring the effects of guanine nucleosides on nucleotides in Jurkat cells using HPLC. We also measured proliferation and cell death with microscopy and fluorescence-activated cell sorting. Guanosine increased GTP to 600% and reduced ATP to 40% of control. This resulted in cell death with a predominance of necrosis. Deoxyguanosine caused similar increases in GTP but at earlier time points. Cell death was severe with a predominance of apoptosis. Deoxyguanosine but not guanosine increased dGTP to 800% of control. Adenosine inhibited the effects of guanosine, in part by competing for uptake. In stimulated leukocytes, guanosine and deoxyguanosine altered the nucleotide pools in a way qualitatively similar to that observed in Jurkat cells. However, proliferation was enhanced rather than impaired. In conclusion, guanosine and deoxyguanosine are toxic to Jurkat cells through two mechanisms: ATP depletion, causing necrosis, and the accumulation of dGTP, resulting in apoptosis.
cancer; lymphocytes; apoptosis; necrosis; adenosine 5'-triphosphate; guanosine 5'-triphosphate
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INTRODUCTION |
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GUANINE NUCLEOTIDE POOLS are four to six times smaller than the adenine nucleotide pools in most mammalian cells (13, 17). Therefore, guanine nucleotides are thought to be more rate limiting for DNA and RNA synthesis than are adenine nucleotides (16). Furthermore, many studies have shown that the guanine nucleotide synthetic machinery is upregulated in various neoplastic cells (18, 30). Enzymes such as inosine monophosphate dehydrogenase (the rate-limiting enzyme for de novo GTP synthesis), GMP synthase, nucleoside diphosphate kinase, and the salvage pathway enzymes all have been shown to have increased activity in malignant cells (31). This finding has led to the successful use of guanine nucleotide synthesis inhibitors in the treatment of various neoplastic disorders (14, 20, 28, 29).
Conversely, an equally large body of data in the cancer literature points to guanine bases and nucleosides as potential antineoplastic agents (19, 22). In fact, many chemically modified guanine bases and nucleosides have been shown to be effective against various types of cancer cells (5, 22). More interestingly, nonmodified guanine bases and nucleosides have also been shown to be toxic to some neoplastic cells (15, 21, 27). These data are paradoxical in the sense that guanine substrates should feed into the salvage pathway and increase cellular GTP levels. Providing additional nucleic acid substrate, in turn, should promote growth and proliferation of the neoplastic cells. Why this does not happen is unclear, but several explanations have been proposed. These include interference with de novo ATP synthesis, direct toxicity of accumulated deoxyribonucleotides, and indirect toxicity through enhancement of the effects of other chemotherapeutic agents (3, 12, 15).
Although each of these explanations is plausible, quantitative data on the effects of purine bases and nucleosides on cellular nucleotide pools and cell function are lacking. In particular, qualitative and quantitative differences in the effects of guanosine and deoxyguanosine on ribonucleotide and deoxyribonucleotide pools have not been examined systematically. Also, the differential effects of guanosine and deoxyguanosine on cell cycle arrest, apoptosis, or necrosis have not been addressed. Thus, although these guanine compounds have some proven usefulness in the treatment of cancer, our knowledge of their effects and mechanism of action is incomplete. It was the aim of this study to investigate the cellular mechanisms by which these compounds act to stop the growth of neoplastic cells.
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METHODS |
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Cell culture. Jurkat cells were maintained in RPMI medium supplemented with 10% fetal bovine serum (FBS) and grown in a humidified atmosphere (95% air-5% CO2) at 37°C. For all experiments, Jurkat cells were plated at a density of 0.5 million cells/ml and used after 24 h of incubation. Human peripheral blood mononuclear cells (PBMC) were isolated by gradient centrifugation of freshly isolated peripheral blood on Histopaque 1077 (Sigma Chemical, St. Louis, MO); erythrocytes were removed using Red-Out (Robbins Scientific, Sunnyvale, CA). PBMC were then washed with PBS and suspended in RPMI medium at a density of 0.5 million cells/ml. For stimulation studies, ionomycin (1 µM) and phorbol 12-myristate 13-acetate (PMA; 10 ng/ml) were used.
Chemicals. Alanosine was a generous gift from the Drug Synthesis and Chemistry Branch of the National Cancer Institute. All other chemicals were from Sigma Chemical unless stated otherwise.
Nucleotide extraction.
Five million cells were used per experimental condition for all HPLC
experiments. At the end of an experimental protocol, cells were
pelleted at 800 g and washed three times with ice-cold PBS.
Extraction was done with 50 µl of ice-cold acetonitrile followed by
150 µl of cold water (1). The soluble and precipitated
fractions were centrifuged at 16,000 g for 10 min at
20°C. The supernatant fraction, kept on ice, was then gassed with
N2 for 30 min to evaporate acetonitrile. The pellet was
solubilized with 1 N NaOH, and the protein content was analyzed by
Coomassie blue assay (Pierce Chemical, Rockford, IL).
Deoxyribonucleotides were obtained by oxidizing the supernatants with
0.5 M NaIO4, 4 M methylamine, and 1 M rhamnose (8).
HPLC. The column used was a 4-µm Nova-Pack C18 cartridge (100 × 8-mm inner diameter) equipped with a radial compression chamber (Waters, Millford, MA). The buffer consisted of 20% acetonitrile, 10 mM ammonium phosphate, and 2 mM PIC-A ion-pairing reagent (Waters) and was run isocratically at 2 ml/min (10). Samples were diluted by half, and the injection volume was 100 µl. An HP Chemstation model 1100 was used (Hewlett-Packard, Wilmington, DE), and the ultraviolet (UV) detector was set at 254 nm. HPLC-grade nucleotide standards were used to calibrate the signals. Internal standards were occasionally added to the samples to test recovery, which exceeded 90% for all nucleotides.
Proliferation. Cell proliferation assessment was done by using Jurkat cells or PBMC in a total of 0.2 ml of medium in sterile 96-well plates. Either 2 h (Jurkat) or 16 h (PBMC) before cells were harvested, 2 mCi of [3H]thymidine (ICN Pharmaceuticals, Costa Mesa, CA) were added to each well. Cells were harvested (Packard Harvester; Packard Instruments, Downers Grove, IL) onto glass fiber filters, and thymidine incorporation was measured with a Beckman liquid scintillation counter. Results were recorded as disintegrations per minute (dpm).
Guanosine uptake. Guanosine uptake was determined by incubating cells with [3H]guanosine (Amersham Pharmacia Biotech, Arlington Heights, IL) for 4 h (Jurkat) or 18 h (PBMC) under the conditions indicated. Cells were centrifuged, washed vigorously with cold PBS, and transferred to scintillation vials. [3H]guanosine uptake was measured with a Beckman liquid scintillation counter. Results were recorded as dpm and are expressed as means ± SE of triplicate samples.
Microscopy. Jurkat cells were plated into 35-mm Corning culture dishes at 0.5 million cells/ml (2 ml/plate). After treatment with an experimental condition, cells were costained with 0.l µg/ml Hoechst 33342 and 1.5 µg/ml propidium iodide for 15 min. Stained cells were centrifuged at 800 g and then resuspended in 16 µl of medium for preparation of a slide smear. A Zeiss confocal microscope (LSM 510), equipped with UV and helium lasers, was used to examine the morphology of the nucleus for apoptotic features such as condensation and fragmentation. Concomitant staining with propidium iodide (yellow) identified necrotic cells.
Fluorescence-activated cell sorting. Cells were plated as described for microscopy but were costained with annexin-V-fluos (Roche, Indianapolis, IN) and propidium iodide per the manufacturer's instructions. Percentages of apoptotic and dead Jurkat cells were determined by two-color immunofluorescence (fluorescence-activated cell sorting, or FACS) using a FACSCalibur flow cytometer (BD Biosciences, San Jose, CA). An acquisition gate was set to include ~30,000 of the centrally located Jurkat cells for each sample acquisition using linear forward scatter vs. linear side scatter. This acquisition strategy resulted in ~50,000 ungated events being included for each sample analysis. Dot-plot integration cursors for the determination of background FL-1 channel (annexin channel) vs. FL-2 channel (propidium iodide channel) fluorescence were set using unstained Jurkat cells. The cursors were placed such that 99% of the unstained cells were contained in the double-negative FL-1 and FL-2 fluorescence quadrant. This integration cursor placement remained unchanged when Jurkat cells stained with annexin and propidium iodide were analyzed.
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RESULTS |
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Effects of guanosine and deoxyguanosine on nucleotides profiles and
proliferation of Jurkat cells.
Jurkat cells exposed to 500 µM guanosine for 24 h showed an
increase in GTP pools to ~600% of control and a decrease in ATP to
~40% of control (Fig. 1, A
and B). Deoxyguanosine increased GTP to ~250% of control
and reduced ATP to ~25% of control (Fig. 1, A and
C). These effects of guanosine and deoxyguanosine were unchanged by the presence of 1 µg/ml mycophenolic acid, an inhibitor of GTP de novo synthesis (data not shown).
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Effects of other guanine and adenine compounds on nucleotides and
proliferation of Jurkat cells.
We next examined whether other bases and nucleosides have effects on
Jurkat cells similar to those seen with guanosine and deoxyguanosine.
As shown in Fig. 4, guanine increased GTP
and reduced ATP and proliferation in a way similar to guanosine.
Adenosine and deoxyadenosine moderately increased ATP to 170% and
150%, respectively, and had no significant effect on GTP and
proliferation. Adenine increased ATP to ~180% of control and caused
only moderate reductions in GTP and proliferation.
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Guanosine and deoxyguanosine and Jurkat cell death.
The profound reductions in proliferation observed with guanosine and
deoxyguanosine indicated either cell cycle arrest with preserved
viability or cell death. To investigate these possibilities, we
examined the cells under confocal microscopy and double nuclear staining (Fig. 5). Control cells showed
normal nuclear morphology and excluded propidium iodide. Occasional
apoptotic nuclei were observed (Fig. 5A). In cells
treated with 500 µM guanosine or deoxyguanosine for 24 h, a
mixed picture of necrosis and apoptosis was observed (Fig. 5,
B and C). Cell death at 24 h was next
quantitated with FACS analysis. Control cells showed 79% viability,
whereas guanosine reduced viability to 54% (Fig.
6, A and B). This
reduction was accounted for by a moderate and equal increase in
apoptosis and necrosis (Fig. 6B). Guanine (but not
adenine or the adenine nucleosides) caused a death/viability profile
similar to that for guanosine (data not shown). Deoxyguanosine exposure
reduced viability to 18%. Although the proportion of necrosis was
similar to that observed with guanosine, close to 50% of cells were
apoptotic, a sixfold increase compared with control and
threefold higher than in guanosine-treated cells (Fig. 6C).
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Mechanisms of cell death. To distinguish between cell death caused by the increase in GTP, the decrease in ATP, or both, we used alanosine, a selective inhibitor of de novo ATP synthesis (25). At 1 mM, alanosine reduced ATP to 40 ± 5% and increased GTP to 115 ± 7% of control. Thus alanosine reduced ATP to levels similar to those observed with the guanine nucleosides but without the exaggerated increase in GTP. As shown in Fig. 6D, alanosine caused cell death with a predominance of necrosis. Therefore, the increase in GTP is not directly required for the induction of a necrotic phenotype.
Whereas ATP depletion appeared to account for the necrosis observed with both nucleosides, it was more difficult to explain the apoptotic response. Both nucleosides caused marked increases in GTP, yet only deoxyguanosine resulted in significant apoptosis. One possibility was that there is a critical GTP threshold above which apoptosis is triggered. However, we have consistently observed a relatively enhanced potency of deoxyguanosine compared with guanosine in reducing proliferation, even with GTP elevations below those reported in Fig. 2 (data not shown). Thus we considered that deoxyguanosine-mediated apoptosis was likely due to effects in addition to those on GTP and ATP. We therefore measured deoxyribonucleotide levels in Jurkat cells exposed to guanosine or deoxyguanosine (Fig. 7). On average, guanosine reduced dGTP to <20% of control and dATP to 90% of control. In contrast, deoxyguanosine caused a dose-related increase in dGTP such that at 500 µM, dGTP levels were ~800% of control. The levels of dATP were not significantly changed. K562 cells showed changes in dATP and dGTP virtually identical to those seen in Jurkat cells (data not shown).
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Effect of adenosine on guanosine uptake and
guanosine- induced changes in nucleotides and proliferation.
With the presumption that the effects of the nucleosides required
cellular uptake, we hypothesized that inhibition of this uptake would
prevent the changes in purine pools and their effect on proliferation.
To test this hypothesis, we studied the effects of the nucleoside
transport inhibitor nitrobenzylthioinosine (NBMPR) (2),
sodium-free medium to inhibit sodium-dependent nucleoside transport
(7), and adenosine as a possible competitive inhibitor (2, 23). Whereas neither 10 mM NBMPR nor sodium-free
medium had any impact on the uptake of [3H]guanosine,
excess adenosine (500 µM vs. 5 µM guanosine) blocked 90% of
guanosine uptake (data not shown). Even at equimolar concentrations (500 µM each), adenosine reduced [3H]guanosine uptake
by 41% at 4 h. This latter finding was accompanied by a reduction
in the increase in GTP and the decrease in ATP observed in the absence
of adenosine (Fig. 8, A and
B). Finally, reduced guanosine uptake was also accompanied
by a decrease in its inhibitory effect on proliferation (Fig.
8C).
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Effects of guanosine and deoxyguanosine on nucleotides and
proliferation of PBMC.
To determine whether the effects of guanosine and deoxyguanosine occur
in normal cells, we investigated their effects on primary human PBMC
(Fig. 9). In unstimulated cells, both
nucleosides caused a moderate increase in GTP levels (<200%) but
without changes in ATP or proliferation. When cells were stimulated
with PMA/ionomycin, guanosine treatment increased proliferation to
300% of control with no change in GTP and a reduction in ATP to 65%
of control. Deoxyguanosine increased GTP and proliferation to 180% and
200% of controls, respectively, and ATP was reduced to 75% of
control. [3H]guanosine uptake was abundant in both
resting and stimulated PBMC (data not shown), indicating that a failure
of toxic effect of guanosine was not due to failure of uptake.
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DISCUSSION |
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Nucleosides and nucleoside analogs have been proposed as useful agents in the treatment of some forms of lymphoid malignancies (9, 21). Their mechanism of action is diverse and probably depends on the specific tumor cell being treated. In this study, we examined nucleotide profile changes induced by guanine nucleosides and their impact on cell death. The lymphoblastic leukemia Jurkat cell line was used as a model for rapidly proliferating neoplastic cells.
A remarkable finding was the exaggerated synthesis of GTP in response to both guanosine and deoxyguanosine. This increased synthesis of up to 800% of control levels indicates a highly upregulated salvage pathway with no feedback inhibition (29-31). The increase in GTP was accompanied by a significant reduction in ATP to 20-40% of control. This decrease in ATP has been reported by others but without measurements of GTP (15). The proposed mechanism for ATP reduction was production of GMP from guanosine and a feedback inhibition by GMP on ATP de novo synthesis. Our measurements of increased GTP provide an additional explanation: ATP could be consumed as the phosphate donor for the conversions of GMP to GDP and GDP to GTP. These reactions are catalyzed by nucleoside diphosphate kinase, an enzyme that equilibrates guanine and adenine nucleotide pools. Inhibitors of this enzyme are not readily available (6, 24, 26), but they would provide direct evidence for the link between ATP depletion and GTP synthesis.
ATP depletion or energy failure was proposed as an important step in chemotherapy-induced tumor regression (3). Our results with alanosine show that ATP depletion without changes in GTP is sufficient to induce death by necrosis. The amount of necrosis seen with guanosine and deoxyguanosine correlated with the degree of ATP depletion they induced. Therefore, a direct toxic effect of increased GTP seems unlikely but cannot be totally excluded. Recently, ATP depletion of varying magnitudes has been linked specifically to the necrotic phenotype in different tissues (4, 11).
While ATP depletion is probably the cause of necrosis, the increased GTP is not likely to be the direct cause of apoptosis. This is because both guanosine and deoxyguanosine caused similar increases in GTP (albeit at different time points) yet induced significantly different amounts of apoptosis. The cause of the mild increase in apoptosis with guanosine is unclear but may be related to the reduction in dGTP induced by guanosine. Deoxyguanosine resulted in large increases in dATP and dGTP, with the latter increasing to 800% of control. These increases in dGTP and dATP may cause apoptosis by inhibiting ribonucleotide reductase. This enzyme converts ribonucleotides to deoxyribonucleotides and offers the only route for the synthesis of deoxyribonucleotides. dGTP is a known potent inhibitor of this enzyme, and increased dGTP levels could therefore result in significant decreases in levels of other deoxyribonucleotides such as dCTP (27). This decrease in other deoxyribonucleotide levels could be the cause of the observed apoptosis. Direct toxicity of increased dGTP or dATP has not been reported but cannot be excluded.
Our results with primary human PBMC and a variety of other cell lines show that the toxicity of guanine bases and nucleosides is not a general phenomenon. However, the finding that rapidly growing K562 human lymphoblast tumor cells show a Jurkat-like pattern of response to guanosine and deoxyguanosine, including potent inhibition of proliferation, indicates that these findings are not limited to the Jurkat cell line. In contrast to K562 cells, resting healthy PBMC were totally unaffected by guanosine or deoxyguanosine, whereas stimulated healthy PBMC showed nucleotide changes qualitatively similar to those in Jurkat cells, with increases in GTP and reductions in ATP. However, these changes were of smaller magnitude and were accompanied by increased proliferation rather than growth arrest and cell death. Whether this difference simply reflects the quantitatively smaller changes in nucleotides in PBMC compared with Jurkat and K562 cells or is a true phenotypic difference is unknown. Nevertheless, it conveys significant selectivity toward neoplastic cells and thus supports the use of these compounds as chemotherapeutic agents.
In summary, we have shown that guanine bases and nucleosides (but not adenine bases and nucleosides) are potent inhibitors of Jurkat cell proliferation. This inhibition was predicated on cellular uptake and metabolism, a process inhibited by adenosine. The inhibition of proliferation resulted from cell cycle arrest at early time points, followed by significant cell death in the form of apoptosis and necrosis. ATP depletion secondary to excessive GTP synthesis correlated best with necrosis. Apoptosis, seen mostly with deoxyguanosine, resulted from exaggerated accumulation of dGTP and dATP. Although the generality of these observations and their applicability to other neoplastic cells remain to be determined, the lack of toxicity to normal primary leukocytes may present an opportunity to influence clinical therapy for some malignancies.
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ACKNOWLEDGEMENTS |
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We are grateful to April Camp for technical assistance.
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FOOTNOTES |
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This work was supported in part by American Cancer Society Grant IRG-84-002-17 (to T. D. Batiuk and P. C. Dagher) and a National Kidney Foundation Clinician-Scientist Award (to T. D. Batiuk).
Address for reprint requests and other correspondence: P. C. Dagher, Division of Nephrology, FH 115, 1120 South Drive, Indianapolis, IN 46202 (E-mail: pdaghe2{at}iupui.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 25 January 2001; accepted in final form 17 July 2001.
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