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Molecular Cancer Research 1:577-588 (2003)
© 2003 American Association for Cancer Research

Cell Death, Cell Cycle and Senescence

Ectopic Expression of 10-Formyltetrahydrofolate Dehydrogenase in A549 Cells Induces G1 Cell Cycle Arrest and Apoptosis1

Natalia V. Oleinik1 and Sergey A. Krupenko1

Department of Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, SC

Requests for reprints: Sergey A. Krupenko, Department of Biochemistry and Molecular Biology, Medical University of South Carolina, 173 Ashley Avenue, Room 512-B BSB, Charleston, SC 29425. Phone: (843) 792-0845; Fax: (843) 792-8565. E-mail: krupenko{at}musc.edu


   Abstract
Top
 Notes
 Abstract
Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
We have recently shown that transient expression of 10-formyltetrahydrofolate dehydrogenase (FDH) strongly inhibits proliferation of several cancer cell lines and ultimately results in cell death. In the present studies using Tet-On system, we have generated a stable A549 lung carcinoma cell line capable of inducible FDH expression. Using this system, we were able to express FDH at different levels depending on concentration of the inducer, doxycycline, and we have observed that inhibition of proliferation depends on FDH intracellular levels. We have further shown that induction of FDH expression results in initiation of apoptosis beginning 24 h post-induction. Apoptotic cells revealed cleavage of poly-(ADP-ribose) polymerase and general caspase inhibitor zVAD-fmk protected cells against FDH-induced apoptosis. FDH-expressing cells showed accumulation of cells in G0-G1 phase and a sharp decrease of cells in S phase. Accumulation of intracellular FDH was followed by accumulation of the tumor suppressor protein p53 and its downstream target p21. These results indicate that FDH antiproliferative effects on A549 cells include both G1 cell cycle arrest and caspase-dependent apoptosis.


   Introduction
Top
 Notes
 Abstract
 Introduction
Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
Folate coenzymes are crucial for cell proliferation due to their involvement in nucleotide biosynthesis and methylation (1–3). Disruption of folate metabolism may produce drastic effects on cells including: altered protein expression (4, 5); decreased DNA repair capability and accumulation of DNA damage (6–12); increased chromosomal aberrations and fragility (13, 14); reduced growth rate (15, 16); and impairment of cell division (1). Because of intensive DNA/RNA synthesis, rapidly proliferating cancer cells have an increased demand for nucleotides (17) and therefore they are highly susceptible to changes in intracellular folate status (18). Prolonged impairment of intracellular folate metabolism induces irreversible metabolic damage and eventually will result in cell death. The important role of folate for cell function is the basis for treatment of tumors by targeting folate metabolism/intracellular folate pools through inhibition of key folate enzymes (19–23). Antifolates, a class of antimetabolite drugs widely used in cancer chemotherapy, inhibit folate-dependent enzymes and exert a powerful impact on folate-mediated cellular processes resulting in inhibition of proliferation and cell death (19–23). Numerous studies in recent years have demonstrated that despite targeting of multiple metabolic pathways, anticancer agents, including antifolates, ultimately kill cells by induction of biochemical cascades associated with apoptosis (24–26). Apoptosis is a primary and universal mechanism that eliminates undesirable cells in response to internal or external signals and it is characterized by distinctive morphological and biochemical changes (26). However, the molecular mechanisms that connect folate metabolism and induction of apoptosis are not clear at present.

The most extensive studies of molecular mechanisms induced by disruption of folate utilizing enzymes were carried out using antimetabolites such as methotrexate (MTX) (27, 28). It has been demonstrated that downstream events include induction of DNA damage (29), cell cycle arrest (30, 31), and apoptosis (31–33). Because MTX, as well as other antifolates, induces both cell cycle arrest and apoptosis in tumor cells, it is likely that disruption of the cell cycle caused by antifolate treatment is a common trigger initiating apoptotic sequences (31). Most antifolates, including inhibitors of de novo purine biosynthesis, kill cells in S phase of the cell cycle (30, 34, 35). Apparently, duplication of the cellular genome in the S phase is a critical event, during which cells are highly susceptible to agents that disturb the tight regulation of synthesis and utilization of DNA precursors (36). It has been also shown that MTX induces expression of tumor suppressor protein, p53 (37). In addition, effects of this drug are mediated by caspases (38–40) while the pro-survival proteins, Bcl-XL and Bcl-2, protect cells against antifolate-induced apoptosis (32, 41). Folate deficiency was also reported to induce apoptosis in both cell culture and animal models (6, 42–44). It has been further concluded that, in HepG2 cell line and erythroblasts, apoptosis induced by folate deficiency is p53 independent (42, 45). Apoptosis induced in the HepG2 cells by media folate deprivation was preceded by G2 cell cycle arrest accompanied by accumulation of cells in S phase of cell cycle (42), in contrast to effects of MTX which arrests cells at G1 (30, 31).

Up-regulation of the folate enzyme, dihydrofolate reductase (DHFR), is a common mechanism of resistance of cancer cells to antifolates (23, 28). Overexpression of DHFR, as well as high levels of some other folate-dependent enzymes, allows supporting favorable rate of nucleotide biosynthesis that promotes proliferation and apparently protects cells against apoptosis. We have recently shown that, in contrast to DHFR, another folate enzyme, 10-formyltetrahydrofolate dehydrogenase (FDH), is down-regulated in tumors and possesses suppressor effects on cancer cells (46). Transient expression of FDH in several cancer cell lines strongly inhibited proliferation and resulted in cell death (46). FDH catalyses conversion of 10-formyltetrahydrofolate (10-formyl-THF) to THF (1). In agreement with the FDH catalytic function, it has been reported that loss of this enzyme in mice increased the 10-formyl-THF and lowered the THF intracellular pools (47). Accordingly, up-regulation of FDH should result in diminished levels of 10-formyl-THF. Importantly, 10-formyl-THF is a substrate for two reactions of de novo purine biosynthesis (48). These reactions are catalyzed by two different enzymes, one of which is glycinamide ribonucleotide formyltransferase (GART) (49). This enzyme has been a target for new antifolates and GART inhibition results in strong suppression of cell proliferation (50). The antiproliferative effects of both GART inhibitors (51) and FDH overexpression (46) can be reversed by supplying enough exogenous purine to maintain intracellular purine pools via salvage pathway. These results imply that antiproliferative effects of FDH are similar to those produced by GART inhibitors and suggest that they might occur through the same molecular mechanism. Compared to mechanisms of MTX, molecular events controlling cell death in response to treatment with GART inhibitors are less clear. Several studies revealed that GART inhibitors result in cell cycle alterations (34, 52) but the question of whether and how they induce apoptosis remains to be elucidated. Interestingly, in contrast to GART inhibitors, antiproliferative effects of FDH appear to be tumor cell specific. Thus, proliferation of two non-tumor origin cell lines, HEK293 (46) and MCF-10A,2 was not affected by FDH expression. Moreover, these cell lines express endogenous FDH in contrast to a number of cancer cell lines which are FDH-deficient (46). Hence, the present study was undertaken to investigate mechanisms of cell death induced by FDH overexpression in stably transfected cells.


   Results
Top
 Notes
 Abstract
 Introduction
 Results
Discussion
 Materials and Methods
 Acknowledgements
 References
 
Generation of Cell Line for Stable FDH Expression
To achieve stable, inducible FDH expression, we have placed FDH under control of a regulated promoter. We employed the Tet-On gene expression system (Clontech, Palo Alto, CA) that is based on use of the Escherichia coli tetracycline-resistance operon (53). The first component of this system, pTet-On regulatory plasmid, allows expression of the regulatory protein. The second component is the response plasmid, which bears the gene of interest, FDH, under control of the tetracycline-response element. The tetracycline-response element is located immediately upstream of the minimal CMV promoter, which lacks strong enhancer elements. Therefore, there is no expression of FDH in the absence of bound regulatory protein. In the presence of doxycycline, a tetracycline derivative, regulatory protein binds to tetracycline-response element and activates transcription of the FDH. Generation of a stable line for inducible FDH expression included two subsequent steps: (a) generation of a cell line stably expressing regulatory protein; and (b) generation of a stable line capable of expressing FDH plus to the regulatory protein (double-stable cells). In the first step, A549 cells were transfected with the pTet-On regulatory plasmid using LipofectAMINE. Transfected clones were selected by resistance to the antibiotic, geniticin (G418). Selected clones that appeared 15 days post-transfection were screened for induction of luciferase expression by transient transfection with the pTRE2hyg-Luc vector. Expression of luciferase in this vector is under control of tetracycline-response element and the vector served as a reporter in this step. Out of a total of 68 clones that have been screened, 5 clones demonstrated 17- to 42-fold induction of luciferase activity by doxycycline. These clones were further transfected with a pTRE2hyg/FDH construct that bears FDH gene and resistance to hygromycin. Hygromycin-resistant clones appeared 29 days post-transfection and they were tested for doxycycline-induced FDH production. Out of a total 80 tested clones, 7 revealed expression of FDH in response to addition of doxycycline and no FDH production was observed in the absence of doxycycline (data not shown).

Characterization of Stable Cell Line Expressing FDH
Two out of seven doxycycline-responsive clones, A549/ATG10.25 and A549/ATG10.26, were used in further studies to evaluate the influence of inducible FDH expression on cellular proliferation. Results obtained using either of these clones were essentially the same. Both clones were generated from the same parental clone, A549/ATG10, which is able to express regulatory protein but not FDH. FDH production was seen beginning 12 h post-induction (not shown) and the highest FDH levels were reached after 2 days (Fig. 1A). We observed that levels of FDH expression depend on doxycycline concentration in the range of 0.5–2.5 µg/ml (Fig. 1A). Further increases in doxycycline concentration to 5.0 µg/ml did not result in increased FDH levels (data not shown). We have also measured FDH activity in cell cytosol extracts at different post-induction times and for different concentrations of doxycycline. These experiments demonstrated that FDH activity reached at induction with 2.5 or 5.0 µg/ml doxycycline were the same and they were more than eight times and about two times higher than levels reached with 0.5 and 1.25 µg/ml doxycycline, correspondingly (Fig. 1B). In general, levels of FDH activity were consistent with FDH protein levels estimated by immunoblot assays (Fig. 1).



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  		FIGURE 1. Inducible FDH expression in a stable Tet-On A549 cell line (clone A549/ATG10.26). A. FDH levels induced by different doxycycline (Dox) concentrations at different post-induction times were evaluated by immunoblot assay with FDH-specific antiserum. B. FDH enzymatic activity measured in cell cytosol extracts at different post-induction times after induction of FDH expression with different doxycycline concentrations: no doxycycline (open circles); 0.5 µg/ml (closed circles); 1.25 µg/ml (open squares); 2.5 µg/ml (closed squares); 5 µg/ml (open triangles). FDH activity was measured as described in "Materials and Methods." Activity is shown in relative units per 105 cells.

 
In accordance with FDH expression levels, different degrees of inhibitory effects on cell growth were observed (Fig. 2A). Levels of FDH achieved at 2.5 or 5 µg/ml doxycycline resulted in cytotoxicity, while induction of FDH expression by 1.25 µg/ml doxycycline resulted in cytostatic effects. And levels of FDH achieved at 0.5 µg/ml doxycycline only slightly delayed cell growth. In control experiments, it was shown that up to 2.5 µg/ml, doxycycline itself did not influence growth of A549 cells (data not shown) or A549/ATG10 cells (Fig. 2B). At 5 µg/ml, it inhibited cell proliferation, but its effects did not appear until day 6 after doxycycline addition (Fig. 2B) and resulted only in cytostatic effects up to 16 days of cell growth (not shown).



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  		FIGURE 2. Effect of stable FDH expression on proliferation of Tet-On A549 (clone A549/ATG10.26) cells. A. Proliferation time course of Tet-On A549 stable cells capable of inducible FDH expression at different doxycycline concentrations: no doxycycline (open circles); 0.5 µg/ml (closed circles); 1.25 µg/ml (open squares); 2.5 µg/ml (closed squares); 5 µg/ml (open triangles). B. Proliferation time course of A549 cells, not capable of FDH expression, at different doxycycline concentrations (doxycycline concentrations are the same as shown for A). Cell viability was assessed by the trypan blue exclusion assay; viable cells (non-stained with trypan blue) were counted using an inverted microscope.

 
We further examined the duration of FDH induction by doxycycline and how that influences cell growth. In contrast to the previous experiments where cells were grown continuously in the presence of doxycycline, in these experiments, the stable cell line was grown in the presence of 2.5 µg/ml doxycycline for 1–6 days. At indicated time periods (Fig. 3), the cell medium was replaced with medium containing no doxycycline. We found that induction with doxycycline for 1 day produces high intracellular FDH and results in cytotoxicity although longer exposure with doxycycline resulted in earlier cell death (Fig. 3). Overall, these experiments confirmed that elevation of FDH results in cytotoxicity in a concentration-dependent manner.



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  		FIGURE 3. Dependence of proliferation of Tet-On A549 stable cells on duration of FDH induction. Proliferation of A549 stable cells after induction with doxycycline for 1 day (open circles); 2 days (close circles); 4 days (open squares); 6 days (closed squares). In all cases, FDH expression was induced by adding 2.5 µg/ml doxycycline into the culture medium. At corresponding times, medium with doxycycline was replaced with regular culture medium containing no doxycycline. Cell viability was assessed by the trypan blue exclusion assay.

 
In the following experiments, we induced FDH expression with 2.5 µg/ml doxycyclin because this concentration of doxycycline did not affect FDH-deficient cells but resulted in strong antiproliferative effects in stable cells capable of FDH expression. Since pulse induction resulted in antiproliferative effects as well as continuous induction, in all our further experiments, we induced FDH by treatment of cells with doxycycline for 2 days.

Study of FDH-Induced Apoptosis
Stable Tet-On A549 cells expressing FDH revealed characteristic morphological changes that were indicative of apoptosis (data not shown). Nuclear condensation observed by staining with Hoechst dye, and DNA fragmentation with appearance of typical DNA ladder (data not shown), also demonstrated that these cells undergo apoptosis. We further evaluated apoptosis in the A549/ATG10.26 stable cell line expressing FDH by using the annexin V labeling assay in combination with propidium iodide (PI) staining, followed by analysis with flow cytometer (54). These experiments showed that FDH expression induces apoptosis in a time-dependent manner (Fig. 4). A significant number of cells became apoptotic 48 h post-induction with a maximum number of apoptotic cells achieved at 72 h (Fig. 5B). The maximum number of apoptotic cells that accumulated 2–4 days post-induction coincided with the highest intracellular FDH levels (Fig. 1). The cells then enter the late apoptotic stage and 5–6 days post-induction, almost the entire cell population becomes dead.



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  		FIGURE 4. Apoptosis detected by fluorescence-activated cell sorting (FACS) flow cytometry in Tet-On A549 (clone A549/ATG10.26) cells. Left panel (Control), cells not induced for FDH expression. Central panel (+Dox), cells induced for FDH expression in the absence of zVAD-fmk. Right panel (+Dox/+zVAD), cells induced for FDH expression in the presence of 50 µM zVAD-fmk. FDH expression was induced by growing cells in the presence of 2.5 µg/ml doxycycline for 2 days. zVAD-fmk was added to media simultaneously with doxycycline. The time after FDH induction is marked on the left. Cells were labeled with FITC-annexin V and PI as described in "Materials and Methods." R3 (double negative), live cells; R4 (annexin V positive, PI negative), early apoptosis; R2 (double positive), late apoptosis; R1 (annexine V negative PI positive), dead cells.

 


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  		FIGURE 5. Distribution of Tet-On A549 (clone A549/ATG10.26) cells between live/apoptotic/dead populations at different times after FDH induction. A. Control cells (no doxycycline added); cells induced for FDH expression by 2.5 µg/ml doxycycline in the absence (B) or presence (C) of zVAD-fmk. Cells were labeled with FITC-annexin V and PI as described in "Materials and Methods" at time periods shown on the plot. FITC-annexin V/PI labeled cells were detected by FACS. Double negative cells were counted as live cells (open bars); annexin V positive, PI negative cells were counted as early apoptotic (gray bars); double positive cells were counted as late apoptotic (crossed bars); and annexin V negative, PI positive cells were counted as dead cells (black bars).

 
We also studied whether or not the pancaspase inhibitor, zVAD-fmk (55), can prevent FDH-induced apoptosis. In our experiments, zVAD-fmk at 50 µM concentration protected cells against FDH-induced cytotoxicity (Fig. 6, A and B). Examination of FDH-expressing cells treated with zVAD-fmk by phase-contrast microscopy showed that their morphology was not changed and was similar to the morphology of the control cells (Fig. 6A). In contrast, FDH-expressing cells, not treated with zVAD-fmk, showed clear morphological changes (Fig. 6A) indicative of dying cells. Annexin V assay of FDH-expressing cells at different post-induction times further revealed that addition of zVAD-fmk significantly reduces, almost to the background level, fraction of cells undergoing apoptosis (Figs. 4 and 5C). There was also no increase in number of dead cells in the presence of zVAD-fmk during the monitoring period (5 days) (Fig. 5C). Moreover, when FDH activity was decreased substantially by day 6 post-induction (Fig. 6C), apparently due to FDH degradation, zVAD-protected cells resume their normal growth rate (Fig. 6B). As a marker of activation of caspases after FDH induction, the cleavage of the nuclear protein poly-(ADP-ribose) polymerase (PARP) was determined (56). We monitored the occurrence of the 86-kDa cleaved product of PARP by immunoblot techniques using anti-PARP specific antibodies. These antibodies recognize both full-length PARP and its cleaved fragment. PARP cleavage became evident at 24 h after treatment (Fig. 6D) indicating that FDH-induced apoptotic cell death involves activation of caspase cascade. Addition of zVAD-fmk completely inhibited cleavage of PARP in these experiments (Fig. 6D) further confirming that FDH-induced apoptosis is caspase dependent.



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  		FIGURE 6. Effect of zVAD-fmk on proliferation of FDH-expressing Tet-On A549 stable cells. A. Phase-contrast microscopy of Tet-On A549 (clone A549/ATG10.26) cells grown in the absence and presence of doxycycline. Left panel (Control), cells grown in the absence of doxycycline (FDH expression is not induced); central panel (+Dox), cells grown in the presence of doxycycline (FDH expression is induced) but in the absence of zVAD-fmk; right panel (+Dox/+zVAD), cells grown in the presence of doxycycline (FDH expression is induced) and in the presence of zVAD-fmk (50 µM). Time post-induction of FDH by doxycycline is shown in the figure. For each sample, a representative area from the middle of cell culture wells is shown. Magnification, x200. B. Proliferation of the cells induced with doxycycline in the absence of zVAD-fmk (open circles) and in the presence of 50 µM zVAD-fmk (closed circles); control (open squares), proliferation of the cells in the absence of doxycycline (uninduced cells). C. Time dependence of FDH activity in doxycycline-induced cells. D. Cleavage of PARP in doxycycline-induced cells in the absence of zVAD-fmk and inhibition of PARP cleavage in the presence of zVAD-fmk. Cell viability was assessed by the MTT assay. PARP (full length and cleaved fragment) was detected by immunoblot with PARP-specific antibodies. FDH expression was induced by growing cells in the presence of 2.5 µg/ml doxycycline for 2 days. zVAD-fmk was added to cells simultaneously with doxycycline.

 
Influence of FDH Expression on Cell Cycle
Using PI staining and flow cytometry (57), we examined DNA content of the stable cell line (clone A549/ATG10.26) after induction of FDH expression in comparison with cells that were not induced. Induction with 2.5 µg/ml doxycyclin was initiated when cells entered log-phase growth. In untreated control cells, there was no significant change in DNA content and cell cycle profiles remained fairly constant throughout the time course (Fig. 7). In contrast, substantial alterations in DNA content of cells expressing FDH were observed (Fig. 7). A drastic decrease of cells containing S phase DNA occurred with this population dropping from 35% to 3.7% 48 h post-induction (Fig. 7). At the same time, a significant increase of cells in G0-G1 phase was observed: from 55% at the beginning of FDH induction to 90% 48 h later (Fig. 7). Depletion of active S phase was further confirmed by labeling cells with BrdUrd and by monitoring of PCNA levels (58). Both incorporation of BrdUrd into DNA and PCNA expression are characteristic of cells synthesizing DNA that occurs only in S phase (58). These experiments revealed sharp decrease of BrdUrd incorporation (Fig. 7) and PCNA levels (data not shown) after FDH induction, indicating decrease in number of cells synthesizing DNA and a complete lack of cells with newly synthesized DNA 48 h post-induction.



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  		FIGURE 7. Alterations in DNA content of Tet-On A549 stable cells expressing FDH. Bivariate distribution of cells between cell cycle phases was evaluated using PI labeling and BrdUrd incorporation at different times after induction of FDH expression. Left panels show DNA content histograms of non-induced cells (Control), cells induced for FDH expression (+Dox), and cells induced for FDH expression in the presence of zVAD-fmk (+Dox/+zVAD). The percentage of cells in different cell cycle phases is shown. Right panels show time-dependent alterations in BrdUrd incorporation of non-induced (Control), induced (+Dox), and induced in the presence of zVAD-fmk (+Dox/+zVAD) cells. R3 (red), G1 phase; R2 (green), S phase; R4 (blue), G2-M phase. The percentage of cells in S phase (cells capable of BrdUrd incorporation) is shown. FDH expression was induced with 2.5 µg/ml doxycycline. zVAD-fmk (50 µM final concentration) was added to cell media simultaneously with doxycycline. At the indicated post-induction time periods, cells were incubated with BrdUrd, fixed, and stained with both FITC-labeled antibodies against BrdUrd and with PI (see "Materials and Methods" for more details).

 
In the presence of zVAD-fmk, distribution of cells between cell cycle phases in doxycycline-induced cells was similar to that observed in the absence of zVAD-fmk. Thus, significant increase cells in G0-G1 phase and decrease cells in S phase was monitored 24–36 h post-induction (Fig. 7). At later times (48 h post-induction) in the presence of zVAD-fmk, G1 arrest was abrogated and distribution of cells within the cell cycle became similar to that of control cells. At that time, significant number of cells traversed G1-S boundary and entered S phase. Incorporation of BrdUrd confirmed active S phase buildup in these cells (Fig. 7).

Accumulation of p53 and p21 in FDH-Expressing Cells
Using immunoblot assays with p53-specific monoclonal antibodies, we have shown that stable FDH expression in A549/ATG10.26 cells was accompanied by an increase in p53 levels (Fig. 8A). As a control, we treated the A549 derivative clone, ATG10 (A549/ATG10), with doxycycline to test whether doxycycline itself affects p53 levels. This clone, in contrast to clones capable of inducible FDH expression, is able to express regulatory protein but not FDH. No increase in p53 levels was observed in these cells after treatment with doxycycline (Fig. 8B). These experiments confirmed that accumulation of p53 is specifically associated with FDH elevation. An important downstream effector in p53-specific growth arrest is p21 (59). Therefore, we also measured levels of p21 in A549/ATG10 cells and A549/ATG10.26 cells induced with doxycycline. This has been done by immunoblot techniques using p21-specific monoclonal antibodies. We observed a significant increase in intracellular p21 levels in A549/ATG10.26 cells overexpressing FDH (Fig. 8A). In contrast, no changes in p21 levels were observed in control cells, A549/ATG10, which do not express FDH (Fig. 8B). To differentiate the p53 effects from the caspase effects, we have examined p53/p21 cellular levels in the FDH-expressing cells treated with zVAD-fmk. We have observed that p53/p21 levels were similar in both zVAD-fmk treated and untreated cells (Fig. 8A).



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  		FIGURE 8. Accumulation of p53 and p21 in FDH-expressing cells. A. Levels of FDH, p53, and p21 in Tet-On A549 stable cells (cells capable to induce FDH expression, clone A549/ATG10.26), at different times after induction with doxycycline in the absence and in the presence of zVAD-fmk. B. Levels of p53 and p21 in A549 stable clones capable of regulatory protein expression but not FDH expression (clone A549/ATG10), in response to doxycycline (control). Doxycycline was added into culture medium at a concentration of 2.5 µg/ml. Levels of FDH, p53, and p21 were detected by immunoblot assays using specific antibodies. Samples were normalized for levels of actin (shown for A).

 

   Discussion
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
Materials and Methods
 Acknowledgements
 References
 
We have recently shown that transient expression of FDH in several FDH-deficient cancer cell lines inhibits proliferation and results in cell death (46). Because of this strong antiproliferative effect, we were unable, using a transient expression system, to study mechanisms that underlay these FDH effects. In the present study using stably transfected cells, we have demonstrated that FDH overexpression results in cell cycle arrest and apoptosis.

Inducible expression in our experiments permitted elimination of FDH antiproliferative effects in uninduced cells. We observed that, similar to experiments with transient FDH expression, stable inducible FDH expression inhibited cell proliferation and resulted in cell death. Using different inducer concentrations, we were able to express FDH at different intracellular levels. These experiments demonstrated that the antiproliferative effects of FDH are strictly dependent on its intracellular concentration and there is a narrow margin between FDH levels that kill cells and those that are not toxic. Thus, at certain FDH concentrations, inhibition of cell proliferation, but not cell death, was observed and at lower FDH levels, there was no effect on cell proliferation. This suggests that the extent of the metabolic changes produced by FDH is important and that there are mechanisms in the cell capable of compensation for some effects induced by FDH. The capacity of the compensatory mechanisms, however, appear to be limited and they cannot overcome higher FDH concentrations. Interestingly, even induction of FDH by short-term exposure to doxycycline resulted in cell death. It is not clear whether this was due to the fact that FDH rapidly induces irreversible metabolic changes in the cell that initiates apoptosis at an early stage or due to constant high intracellular FDH levels that persist for several days after short-term induction. Because the half-life for FDH is not known, it is also not clear mechanistically how such high FDH levels occur. It could be due to a long FDH half-life or due to continuous transcriptional induction because of accumulation of the inducer, doxycycline, within the cell. Nevertheless, our experiments showed that after short-term induction, levels of FDH stayed relatively constant through 4 days post-induction. Continuous induction of FDH expression resulted in faster cell death: about 20% of cells were still viable 8 days post-induction in the case of 1-day exposure to doxycycline. On the other hand, there were no live cells by day 7 in the case of continuous induction. This indicates that a decrease of FDH levels has a positive impact on cell survival.

In the present studies, we have demonstrated that FDH cytotoxic effects are mediated through apoptotic pathways. Furthermore, the fact that the pancaspase inhibitor, zVAD-fmk (55) blocked FDH-induced apoptosis, implicates involvement of a caspase cascade mechanism in FDH cytotoxic effects. Cleavage of PARP, a substrate for several caspases, including the most prevalent caspase-3 (60), in FDH-expressing cells further supported a role for caspases in FDH-induced apoptosis. Likewise, as would be expected, cleavage of PARP was blocked in zVAD-fmk-protected cells. The kinetics of proliferation of cells protected with zVAD-fmk is very interesting. Up to day 4 post-induction, when FDH activity is high, cellular proliferation is inhibited but cells do not die. Apparently, zVAD-fmk allows cells to prolong cell cycle arrest without entering apoptosis until the moment when FDH was cleared from the cells. In fact, between day 4 and day 6 post-induction, when FDH activity has decreased sharply (Fig. 6C), cells were released from G1 arrest and resume their normal growth rate. G1 arrest takes place in FDH-expressing cells before they undergo apoptosis. Both accumulation of cells in G0-G1 phase and a rapid decrease of the number of cells in S phase imply that cells did not progress through G1 into S phase while cells in S phase continued to progress to G2-M. Cell cycle arrest in these cells occurred soon after FDH induction and was followed by apoptosis that became evident 12–24 h later. However, PARP cleavage observed as early as 24 h post-induction suggests that some apoptotic events were initiated simultaneously with cell cycle arrest.

Cells activate checkpoint pathways that delay progression through the cell cycle in response to DNA damage to allow time to repair it (61). If DNA repair fails, apoptosis will be triggered (62). Intracellular dNTP pool is crucial for DNA biosynthesis and repair. The FDH substrate, 10-formyl-THF, is required for two reactions of de novo purine biosynthesis (48). We observed that FDH expression in deficient cells decreases 10-formyl-THF more than twice.2 This is expected to decrease the rate of de novo purine biosynthesis with the ultimate depletion of intracellular purine pools. The size of the nuclear dNTP pool is only sufficient to support DNA synthesis for 0.5 to 3 min (43). Therefore, impairment of DNA/RNA synthesis and DNA repair is likely to be the most rapid response to intracellular FDH elevation. As a result, inhibition of cell proliferation and accumulation of DNA damage are expected. In many cases, G1 checkpoint is controlled by p53 tumor suppressor, a transcription factor that is also one of the major regulators of apoptosis (59, 61). DNA-interactive drugs and {gamma} radiation, factors that cause DNA damage directly, generate a p53-dependent G1 arrest (63). Altered nucleotide pools can be also a signal for activation of the p53 pathway (64). p53-mediated transcriptional activation of p21, an inhibitor of cyclin-dependent kinases (61, 65), is the major pathway of p53-induced G1 arrest (66). Studies on p21-null mice have demonstrated that p21 is required to activate G1 cell cycle arrest in response to DNA damage induced by nucleotide pool disruption (67). The p53 gene is inactivated by mutations in over 50% of human cancers (68). However, A549 cells have a wild-type p53 (52) and it would be expected that they have a functional cell cycle checkpoint under control of p53 protein. Thus, G1 arrest in FDH-induced cells accompanied by subsequent accumulation of p53 and its downstream target p21 suggests a role for p53-dependent pathways in FDH antiproliferative effects.

It is not clear at present, however, whether p53 is also involved in FDH-induced apoptosis. Our previous studies on transient FDH expression (46) demonstrated that FDH elevation in deficient cells results in cytotoxicity to cell lines possessing wild-type or mutant p53 as well as in p53-null cells. Furthermore, because p21 in most cases is a negative modulator of apoptosis (69), its up-regulation following accumulation of p53 suggests that while in FDH-induced cells G1 arrest apparently is mediated by p53, apoptosis occurs through a different mechanism. Many studies indicate that p21 protein normally protects cells from p53-induced apoptosis by holding them in cell cycle arrest (see Refs. 66 and 69 and references within). Alternatively, failure to express p21 can lead to apoptotic cell death in response to genotoxic stress (70, 71). One of the possible mechanisms for p21 protection from apoptosis is to bind to procaspases, including procaspase-3, preventing cleavage to form active caspases (72). However, in our experiments, activation of p21 did not prevent cleavage of PARP, a substrate of several caspases including caspase-3, suggesting that this mechanism did not take place in our model. While p21 can protect procaspases from proteolytic activation, it is unable to inhibit already activated caspases (72). PARP cleavage and p21 elevation happened concurrently after FDH induction that explains why apoptosis can still take place despite induction of p21. Although p21 can prevent caspase activation at the same time, the protein itself is a caspase substrate (69). Cleavage of p21 by caspase-3 could be a mechanism to avoid p21-mediated protection against apoptosis. In our study, when apoptosis was blocked with the caspase-3 inhibitor zVAD-fmk, levels of p21 were unchanged. This suggests that it is unlikely that p21 cleavage by caspase-3 plays a role in mechanisms of FDH-induced apoptosis. Furthermore, p53 accumulation and G1 arrest were observed in cells that undergo apoptosis as well as in cells protected from apoptosis with zVAD-fmk. This implies that in FDH-induced cells, G1 arrest and apoptosis happen concurrently rather than sequentially and they might proceed through independent mechanisms. Thus, whereas FDH-induced G1 cell cycle arrest most likely is p53 dependent, FDH-induced apoptosis might involve p53-independent pathways. This would not be entirely unexpected. Indeed, p53-independent apoptosis was observed in A549 cells under circumstances of p53 accumulation and G1 cell cycle arrest (73). Moreover, it has been reported that expression of exogenous wild-type p53 in A549 cells does not induce apoptosis (74), suggesting that, in this cell line, p53-mediated apoptotic mechanisms are impaired.

In contrast to some other key enzymes of folate metabolism, such as DHFR or GART, up-regulation of which is favorable for cell proliferation (17, 23, 28), FDH overexpression works in the opposite direction. In this regard, FDH elevation would be expected to produce an impact on cells similar to that of antifolates targeting GART. Both FDH and GART inhibitors presumably target de novo purine biosynthesis. It is interesting, however, that FDH displays antiproliferative effects that are different from effects of the GART inhibitor AG2034 (52). It has been reported that this inhibitor suppresses proliferation but does not kill A549 cells (52), while FDH overexpression resulted in death of these cells. To explain the effect of the GART inhibitor, it has been proposed that in wild-type p53 cells, with a functional G1 checkpoint, such as A549 cells, G1 arrest occurs in response to purine pool depletion (52). Arrested cells remain viable while cells that have already traversed the checkpoint into S phase die. At present, it is not clear why, in contrast to cells treated with the GART inhibitor AG2034, cells arrested at G1 phase in response to FDH overexpression undergo rapid apoptosis. One explanation could be that because FDH continuously recycles 10-formyl-THF, DNA damage will be eventually accumulated beyond repair limit resulting in a signal to undergo apoptosis. Furthermore, it is well known that even with drugs that target the same metabolic pathway, the sequence of events preceding apoptotic cell death as well as apoptotic mechanisms can be different (31). In fact, other antifolates, including inhibitors of GART, revealed either no dependence of inhibitory effects on p53 status or stronger inhibitory effect on cells expressing wild-type p53 (75–77), as well as an absence of induction of G1 arrest (77). It should be also mentioned that we did not evaluate GART activity in our study. Therefore, we cannot rule out the possibility that FDH-induced cell death might be independent of reduction of GART function.


   Materials and Methods
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
Acknowledgements
 References
 
Cell Culture
Cell media and reagents were purchased from Invitrogen, Inc. (Carlsbad, CA) unless otherwise indicated. Human A549 lung non-small carcinoma cells were obtained from American Type Culture Collection. Cells were grown at 37°C in a humidified atmosphere containing 5% CO2 in RPMI 1640 supplemented with 10% fetal bovine serum (Atlanta Biologicals, Atlanta, GA), L-glutamine (2 mM), and sodium pyruvate (1 mM). A549 stable clones were grown on the same media containing Tet-certified serum (10%) instead of regular serum, and corresponding antibiotics. FDH expression was induced by doxycycline. Antibiotics and Tet-certified serum were purchased from Clontech. Cell viability was assessed by trypan blue exclusion or by MTT cell proliferation assay as described elsewhere (58). MTT cell proliferation assay was performed using CellTiter 96 kit (Promega, Madison, WI) according to the manufacturer's directions.

Vector Generation for Stable FDH Expression
A fragment that included the entire coding sequence for FDH plus an extra 700 bp of pcDNA3.1 on the 5' end was excised from the pcDNA3.1/FDH construct (46) using MluI and NotI restriction endonucleases and cloned into the pTRE2hyg vector (Clontech) through corresponding restriction sites. The MluI site was than converted to AflII site using site-directed mutagenesis with Quick Change kit (Stratagene, La Jolla, CA). The second AflII restriction site was originally present in the insert immediately upstream of the FDH coding sequence. The extra non-coding part of the insert (700 bp) was excised with the AflII restriction endonuclease with the subsequent ligation carried out using Fast Ligation kit (Roche Molecular Biochemicals, Indianapolis, IN).

Generation of Stable Tet-On Cell Lines
Cells at about 70% confluence were transfected with 4 µg Tet-On vector (Clontech) using LipofectAMINE Plus Reagent (Invitrogen) according to the manufacturer's protocol. After transfection, cells were plated in five 10-cm-diameter cell culture dishes at 2.0 x 105 cells/plate density. Transfected cells were selected by resistance to antibiotic G418. Antibiotic was added 48 h post-transfection at a concentration of 700 µg/ml. In preliminary experiments, it has been found that this concentration is sufficient to prevent growth of non-transfected cells. Appearance of geneticin resistant clones was monitored using an inverted microscope. These clones were isolated using cloning cylinders, transferred to individual wells, and screened for induction of luciferase activity. Experiments were performed in six-well plates. Cells at about 70% confluence were transfected with 2 µg of pTRE-Luc vector using LipofectAMINE. Doxycycline was added at a concentration of 1.5 µg/ml. Forty-eight hours post-transfection, cells were collected and luciferase activity was assayed using the Luciferase Reporter Assay Kit (Clontech).

Generation of a Double-Stable Cell Line for Inducible FDH Expression
Tet-On A549 cells were grown to about 70% confluence and transfected with the pTRE2hyg/FDH vector construct using LipofectAMINE Plus reagent. Forty-eight hours later, hygromycin was added to the medium at a concentration of 400 µg/ml. The appearance of positive transfectants (hygromycin resistant cells) was monitored visually using an inverted microscope. Hygromycin resistant clones were further examined for induction of FDH expression. These clones were grown to about 50% confluence and doxycycline was added to a final concentration of 0.5–5.0 µg/ml. Intracellular FDH was measured in cell lysates 48 h later using immunoblot techniques.

Immunoblot Analysis
Attached cells (about 1 x 106) were washed free of culture medium by rinsing in PBS and lysed by adding 50 mM Tris-HCl buffer (pH 8.0) containing 0.15 M NaCl, 2 mM EDTA, 1% Triton X-100 and protease inhibitors (Sigma, St. Louis, MO). Cell lysates were subjected to SDS-PAGE followed by immunobloting with the corresponding antibodies. FDH was determined in the cell lysate by immunobloting with FDH-specific polyclonal antiserum (1/1500 dilution) (78); p53, p21, and actin were detected using monoclonal antibodies (1/200, 1/100, and 1/5000 dilution, correspondingly) (Oncogene, San Diego, CA); PARP was detected using monoclonal antibodies (1/2000 dilution) from Biomol Research Laboratories, Inc. (Plymouth Meeting, PA). In all cases, immunobloting procedures were carried out using an ECL kit and Hybon-C nitrocellulose membranes (both from Amersham Pharmacia Biotech, Piscataway, NJ) according to the manufacturer's protocol.

Assay of FDH Activity
FDH activity was measured in cell cytosol extracts obtained essentially as described (79). Briefly, cells grown in 25-cm2 flasks were rinsed twice with PBS and resuspended in ice-cold buffer (50 mM Tris-HCl, 100 mM 2-ME, and protease inhibitor cocktail) using a plastic cell scraper. Cell suspensions were sonicated using a Fisher Scientific Model 500 Ultrasonic Dismembrator (five 5-s, 30-W pulses with 1-min intervals of cooling on ice) and insoluble material was precipitated by centrifugation (14,000xg for 5 min at 4°C). Samples were adjusted to yield 1 ml of cytosol from about 2 x 106 cells in each case. All assays were performed at 30°C in a Shimadzu 2401PC double beam spectrophotometer. The reaction mixture contained 0.05 M Tris-HCl (pH 7.8), 100 mM 2-ME, 100 µC NADP+, and 5 µv of substrate, 10-formyl-DDF. The reaction was started by the addition of cytosol (50 µl) in a final volume of 1.0 ml and read against a blank cuvette containing all components except cytosol. Appearance of product, 5,8-dideazafolate, was measured at 295 nm using a molar extinction coefficient of 18.9 x 103 (78).

Annexin V Labeling
Cells were labeled with annexin V and PI using an Annexing-V-FLUOS Staining Kit (Roche Molecular Biochemicals). Experiments were performed according to the manufacturer's protocol. All cells (attached and floating) were used in these experiments. Annexin V and PI staining was detected by cell flow cytometry.

Cell Cycle Analysis
Cells (about 6 x 106) were labeled with BrdUrd for 45 min using the In Situ Proliferation Kit, FLUOS (Roche Molecular Biochemicals). After that, cells were rinsed twice with PBS, detached by trypsin treatment, and fixed with mixture of absolute ethanol and 50 mM glycine (pH 2.0) at a ratio of 7:3, overnight at 4°C. Fixed cells were denatured with 4 M HCl for 20 min at room temperature and incubated in blocking solution for 10 min to prevent non-specific binding. Cells were incubated with anti-BrdUrd antibodies for 45 min at 37°C in a humid chamber. Then the sample was labeled with PI using Cellular DNA Flow Cytometric Analysis Kit (Roche Molecular Biochemicals) according to the manufacturer's protocol. BrdUrd and PI staining was analyzed by cell flow cytometry. PCNA was measured as described (80) using FITC-conjugated monoclonal anti-PCNA antibodies pc10 (BD Biosciences PharMingen, San Diego, CA).

Flow Cytometry
Flow cytometric analysis was carried out in the Hollings Cancer Center core facility on a Becton Dickinson FACSCalibur. Data analysis was performed using CellQuest and Mod Fit software (Becton Dickinson, Palo Alto, CA).


   Acknowledgements
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
References
 
The authors thank Dr. David G. Priest for helpful discussion and critical reading of the manuscript.


   Notes
Top
 Notes
Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
1 NIH grant DK54388. Back

2 S. Krupenko and N. Oleinik, unpublished observation. Back

Received October 2, 2002; revised April 16, 2003; accepted April 17, 2003.


   References
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 

  1. Wagner, C. Biochemical role of folate in cellular metabolism. In: L. B. Bailey (ed.), Folate in Health and Disease, pp. 23–42. New York, NY: Marcel Dekker, Inc., 1995.
  2. Bailey, L. B. and Gregory, J. F. Folate metabolism and requirements. J. Nutr., 129: 779–782, 1999.[Abstract/Free Full Text]
  3. Friso, S. and Choi, S. W. Gene-nutrient interactions and DNA methylation. J. Nutr., 132: 2382S–2387S, 2002.[Abstract/Free Full Text]
  4. Ryan, B. M. and Weir, D. G. Relevance of folate metabolism in the pathogenesis of colorectal cancer. J. Lab. Clin. Med., 138: 164–176, 2001.[Medline]
  5. Jhaveri, M. S., Wagner, C., and Trepel, J. B. Impact of extracellular folate levels on global gene expression. Mol. Pharmacol., 60: 1288–1295, 2001.[Abstract/Free Full Text]
  6. James, S. J., Basnakian, A. G., and Miller, B. J. In vitro folate deficiency induces deoxynucleotide pool imbalance, apoptosis, and mutagenesis in Chinese hamster ovary cells. Cancer Res., 54: 5075–5080, 1994.[Abstract/Free Full Text]
  7. Blount, B. C., Mack, M. M., Wehr, C. M., MacGregor, J. T., Hiatt, R. A., Wang, G., Wickramasinghe, S. N., Everson, R. B., and Ames, B. N. Folate deficiency causes uracil misincorporation into human DNA and chromosome breakage: implications for cancer and neuronal damage. Proc. Natl. Acad. Sci. USA, 94: 3290–3295, 1997.[Abstract/Free Full Text]
  8. Choi, S. W. and Mason, J. B. Folate and colorectal carcinogenesis: is DNA repair the missing link? Am. J. Gastroenterol., 93: 2013–2016, 1998.[Medline]
  9. Duthie, S. J. and Hawdon, A. DNA instability (strand breakage, uracil misincorporation, and defective repair) is increased by folic acid depletion in human lymphocytes in vitro. FASEB J., 12: 1491–1497, 1998.[Abstract/Free Full Text]
  10. Kruman, I. I., Kumaravel, T. S., Lohani, A., Pedersen, W. A., Cutler, R. G., Kruman, Y., Haughey, N., Lee, J., Evans, M., and Mattson, M. P. Folic acid deficiency and homocysteine impair DNA repair in hippocampal neurons and sensitize them to amyloid toxicity in experimental models of Alzheimer's disease. J. Neurosci., 22: 1752–1762, 2002.[Abstract/Free Full Text]
  11. Duthie, S. J., Narayanan, S., Blum, S., Pirie, L., and Brand, G. M. Folate deficiency in vitro induces uracil misincorporation and DNA hypomethylation and inhibits DNA excision repair in immortalized normal human colon epithelial cells. Nutr. Cancer, 37: 245–251, 2000.[Medline]
  12. Duthie, S. J. Folic acid deficiency and cancer: mechanisms of DNA instability. Br. Med. Bull., 55: 578–592, 1999.[Abstract/Free Full Text]
  13. Reidy, J. A. Role of deoxyuridine incorporation and DNA repair in the expression of human chromosomal fragile sites. Mutat. Res., 200: 215–220, 1988.[Medline]
  14. Glover, T. W. Instability at chromosomal fragile sites. Recent Results. Cancer Res., 154: 185–199, 1998.
  15. Borman, L. S. and Branda, R. F. Nutritional folate deficiency in Chinese hamster ovary cells. I. Characterization of the pleiotropic response and its modulation by nucleic acid precursors. J. Cell. Physiol., 140: 335–343, 1989.[Medline]
  16. Huang, R-F. S., Ho, Y-H., Lin, H-L., Wei, J-S., and Liu, T-S. Folate deficiency induces a cell cycle-specific apoptosis in HepG2 cells. J. Nutr., 29: 25–31, 1999.
  17. Weber, G. Biochemical strategy of cancer cells and the design of chemotherapy: G.H.A. Clowes Memorial Lecture. Cancer Res., 43: 3466–3492, 1983.[Free Full Text]
  18. Kim, Y-I. Folate and carcinogenesis: evidence, mechanisms, and implications. J. Nutr. Biochem., 10: 66–88, 1999.[Medline]
  19. Schweitzer, B. I., Dicker, A. P., and Bertino, J. R. Dihydrofolate reductase as a therapeutic target. FASEB J., 4: 2441–2452, 1990.[Abstract]
  20. Priest, D. G. and Bunni, M. A. Folates and folate antagonists in cancer chemotherapy. In: L. B. Bailey (ed.), Folate in Health and Disease, pp. 379–403. New York, NY: Marcel Dekker, Inc., 1995.
  21. Jackson, R. C. Antifolate drugs. In: A. L. Jackman (ed.), Antifolate Drugs in Cancer Therapy, pp. 1–12. Totowa, NJ: Humana Press Inc., 1999.
  22. Kaye, S. B. New antimetabolites in cancer chemotherapy and their clinical impact. Br. J. Cancer, 78: 1–7, 1998.
  23. Banerjee, D., Mayer-Kuckuk, P., Capiaux, G., Budak-Alpdogan, T., Gorlick, R., and Bertino, J. R. Novel aspects of resistance to drugs targeted to dihydrofolate reductase and thymidylate synthase. Biochim. Biophys. Acta, 1587: 164–173, 2002.[Medline]
  24. Thompson, C. B. Apoptosis in the pathogenesis and treatment of disease. Science, 267: 1456–1462, 1995.[Abstract/Free Full Text]
  25. Martin, D. S., Bertino, J. R., and Koutcher, J. A. ATP depletion + pyrimidine depletion can markedly enhance cancer therapy: fresh insight for a new approach. Cancer Res., 60: 6776–6783, 2000.[Free Full Text]
  26. Israels, L. G. and Israels, E. D. Apoptosis. Stem Cells, 17: 306–313, 1999.[Abstract/Free Full Text]
  27. Van Triest, B., Pinedo, H. M., Giaccone, G., and Peters, G. J. Downstream molecular determinants of response to 5-fluorouracil and antifolate thymidylate synthase inhibitors. Ann. Oncol., 11: 385–391, 2000.[Abstract/Free Full Text]
  28. Chu, E. and Allegra, C. J. Antifolates. In: B. A. Chabner and D. L. Longo (eds.), Cancer Chemotherapy and Biotherapy: Principles and Practice, pp. 109–148. Philadelphia, PA: Lippincott-Raven Publishers, 1996.
  29. Kinsella, A. R., Smith, D., and Pickard, M. Resistance to chemotherapeutic antimetabolites: a function of salvage pathway involvement and cellular response to DNA damage. Br. J. Cancer, 75: 935–945, 1997.[Medline]
  30. Tsurusawa, M., Niwa, M., Katano, N., and Fujimoto, T. Flow cytometric analysis by bromodeoxyuridine/DNA assay of cell cycle perturbation of methotrexate-treated mouse L1210 leukemia cells. Cancer Res., 48: 4288–4293, 1988.[Abstract/Free Full Text]
  31. Huschtscha, L. I., Bartier, W. A., Andersson Ross, C. E., and Tattersall, M. H. N. Characteristics of cancer cell death after exposure to cytotoxic drugs in vitro. Br. J. Cancer, 73: 54–60, 1996.[Medline]
  32. Miyashita, T. and Reed, J. C. Bcl-2 oncoprotein blocks chemotherapy-induced apoptosis in a human leukemia cell line. Blood, 81: 151–157, 1993.[Abstract/Free Full Text]
  33. Genestier, L., Paillot, R., Fournel, S., Ferraro, C., Miossec, P., and Revillard, J. P. Immunosuppressive properties of methotrexate: apoptosis and clonal deletion of activated peripheral T cells. J. Clin. Invest., 102: 322–328, 1998.[Medline]
  34. Tonkinson, J. L., Marder, P., Andis, S. L., Schultz, R. M., Gossett, L. S., Shin, C., and Mendelsohn, L. G. Cell cycle effects of antifolate antimetabolite: implications for cytotoxicity and cytostasis. Cancer Chemother. Pharmacol., 39: 521–531, 1997.[Medline]
  35. Seki, K., Yoshikawa, H., Shiiki, K., Hamada, Y., Akamatsu, N., and Tasaka, K. Cisplatin (CDDP) specifically induces apoptosis via sequential activation of caspase-8, -3 and -6 in osteosarcoma. Cancer Chemother. Pharmacol., 45: 199–206, 2000.[Medline]
  36. Hatse, S., De Clercq, E., and Balzarini, J. Role of antimetabolites of purine and pyrimidine nucleotide metabolism in tumor cell differentiation. Biochem. Pharmacol., 58: 539–555, 1999.[Medline]
  37. Heenen, M., Laporte, M., Noel, J. C., and de Graef, C. Methotrexate induces apoptotic cell death in human keratinocytes. Arch. Dermatol. Res., 290: 240–245, 1998.[Medline]
  38. Los, M., Herr, I., Friesen, C., Fulda, S., Schulze-Osthoff, K., and Debatin, K. M. Cross-resistance of CD95- and drug-induced apoptosis as a consequence of deficient activation of caspases (ICE/Ced-3 proteases). Blood, 90: 3118–3129, 1997.[Abstract/Free Full Text]
  39. Cory, A. H., Hickerson, D. H., and Cory, J. G. Apoptosis induced by inhibitors of nucleotide synthesis in deoxyadenosine-resistant leukemia L1210 cells that lack p53 expression. Anticancer Res., 20: 4171–4178, 2000.[Medline]
  40. Papaconstantinou, H. T., Xie, C., Zhang, W., Ansari, N. H., Hellmich, M. R., Townsend, C. M. Jr., and Ko, T. C. The role of caspases in methotrexate-induced gastrointestinal toxicity. Surgery, 130: 859–865, 2001.[Medline]
  41. Simonian, P. L., Grillot, D. A., and Nunez, G. Bcl-2 and Bcl-XL can differentially block chemotherapy-induced cell death. Blood, 90: 1208–1216, 1997.[Abstract/Free Full Text]
  42. Huang, R-F. S., Ho, Y-H., Lin, H-L., Wei, J-S., and Liu, T-S. Folate deficiency induces a cell cycle-specific apoptosis in HepG2 cells. J. Nutr., 29: 25–31, 1999.
  43. James, S. J., Miller, B. J., Basnakian, A. G., Pogribny, I. P., Pogtibna, M., and Muskhelishvili, L. Apoptosis and proliferation under conditions of deoxynucleotide pool imbalance in liver of folate/methyl deficient rats. Carcinogenesis, 18: 287–293, 1997.[Abstract/Free Full Text]
  44. Koury, M. J. and Horne, D. W. Apoptosis mediates and thymidine prevents erythroblast destruction in folate deficiency anemia. Proc. Natl. Acad. Sci. USA, 91: 4067–4071, 1994.[Abstract/Free Full Text]
  45. Koury, M. J., Price, J. O., and Hicks, G. G. Apoptosis in megaloblastic anemia occurs during DNA synthesis by a p53-independent, nucleoside-reversible mechanism. Blood, 96: 3249–3255, 2000.[Abstract/Free Full Text]
  46. Krupenko, S. A. and Oleinik, N. V. FDH, one of the major folate enzymes, is down-regulated in tumor tissues and possesses suppressor effects on cancer cells. Cell Growth & Differ., 5: 227–236, 2002.
  47. Champion, K. M., Cook, R. J., Tollaksen, S. L., and Giometti, C. S. Identification of a heritable deficiency of the folate-dependent enzyme 10-formyltetrahydrofolate dehydrogenase in mice. Proc. Natl. Acad. Sci. USA, 91: 11338–11342, 1994.[Abstract/Free Full Text]
  48. Benkovic, S. J. The transformylase enzymes in de novo purine biosynthesis. Trends Biochem. Sci., 9: 320–322, 1984.
  49. Daubner, S. C., Schrimsher, J. L., Schendel, F. J., Young, M., Henikoff, S., Patterson, D., Stubbe, J., and Benkovic, S. J. A multifunctional protein possessing glycinamide ribonucleotide synthetase, glycinamide ribonucleotide transformylase, and aminoimidazole ribonucleotide synthetase activities in de novo purine biosynthesis. Biochemistry, 24: 7059–7062, 1985.[Medline]
  50. Beardsley, G. P., Moroson, B. A., Taylor, E. C., and Moran, R. G. A new folate antimetabolite, 5,10-dideaza-5,6,7,8-tetrahydrofolate is a potent inhibitor of the de novo purine synthesis. J. Biol. Chem., 264: 328–333, 1989.[Abstract/Free Full Text]
  51. Pizzorno, G., Moroson, B. A., Cashmore, A. R., and Beardsley, G. P. Effects of 5,10-dideaza-5,6,7,8-tetrahydrofolate on nucleotide metabolism in CCRF-CEM cells. Cancer Res., 51: 2291–2295, 1991.[Abstract/Free Full Text]
  52. Zhang, C. C., Boritzki, T. J., and Jackson, R. C. An inhibitor of glycinamide ribonucleotide formyltransferase is selectively cytotoxic to cells that lack a functional G1 checkpoint. Cancer Chemother. Pharmacol., 41: 223–228, 1998.[Medline]
  53. Paillard, F. "Tet-on": a gene switch for the exogenous regulation of transgene expression. Hum. Gene Ther., 9: 983–985, 1998.[Medline]
  54. Darzykiewicz, Z., Bedner, E., and Li, X. Analysis of cell death by flow and laser-scanning cytometry. In: G. P. Studzinski (ed.), Apoptosis. A Practical Approach, pp. 57–80. New York, NY: Oxford University Press, 1999.
  55. Pronk, G. J., Ramer, K., Amiri, P., and Williams, L. T. Requirement of an ICE-like protease for induction of apoptosis and ceramide generation by REAPER. Science, 271: 808–810, 1996.[Abstract]
  56. Kaufman, S. H., Desnoyers, S., Ottaviano, Y., and Poirier, G. G. Specific proteolytic cleavage of poly (ADP-ribose) polymerase: an early marker of chemotherapy-induced apoptosis. Cancer Res., 53: 3976–3985, 1993.[Abstract/Free Full Text]
  57. Crissman, H. A. Cell cycle analysis by flow cytometry. In: G. P. Studzinski (ed.), Cell Growth and Apoptosis. A Practical Approach, pp. 21–43. New York, NY: Oxford University Press, 1995.
  58. Wieder, R. Selection of methods for measuring proliferation. In: G. P. Studzinski (ed.), Cell Growth, Differentiation and Senescence. A Practical Approach, pp. 1–60. New York, NY: Oxford University Press, 1999.
  59. Levine, A. J. p53, the cellular gatekeeper for growth and division. Cell, 88: 323–331, 1997.[Medline]
  60. Tewari, M., Quan, L. T., O'Rourke, K., Desnoyers, S., Zeng, Z., Beidler, D. R., Poirier, G. G., Salvesen, G. S., and Dixit, V. M. Yama/CPP32 ß, a mammalian homolog of CED-3, is a CrmA-inhibitable protease that cleaves the death substrate poly (ADP-ribose) polymerase. Cell, 81: 801–809, 1995.[Medline]
  61. O'Connor, P. M. Mammalian G1 and G2 phase checkpoints. Cancer Surv., 29: 151–182, 1997.[Medline]
  62. Norbury, C. J. and Hickson, I. D. Cellular response to DNA damage. Annu. Rev. Biochem., 41: 367–401, 2001.
  63. Kastan, M. B., Onyekwere, O., Sidransky, D., Vogelstein, B., and Craig, R. W. Participation of p53 protein in the cellular response to DNA damage. Cancer Res., 51: 6304–6311, 1991.[Medline]
  64. Linke, S. P., Clarkin, K. C., Di Leonardo, A., Tsou, A., and Wahl G. M. A reversible, p53-dependent G0/G1 cell cycle arrest induced by ribonucleotide depletion in the absence of detectable DNA damage. Genes Dev., 10: 934–947, 1996.[Abstract/Free Full Text]
  65. Vogelstein, B., Lane, D., and Levine, A. J. Surfing the p53 network. Nature, 408: 307–310, 2000.[Medline]
  66. Roninson, I. B. Oncogenic functions of tumour suppressor p21(Waf1/Cip1/Sdi1): association with cell senescence and tumour-promoting activities of stromal fibroblasts. Cancer Lett., 171: 1–14, 2002.
  67. Deng, C., Zhang, P., Harper, J.W., Elledge, S.J., and Leder, P. Mice lacking p21CIP1/WAF1 undergo normal development, but are defective in G1 checkpoint control. Cell, 82: 675–684, 1995.[Medline]
  68. Zornig, M., Hueber, A., Baum, W., and Evan. G. Apoptosis regulators and their role in tumorigenesis. Biochim. Biophys. Acta, 1551: F1–F37, 2001.[Medline]
  69. Dotto, G. P. p21(WAF1/Cip1): more than a break to the cell cycle? Biochim. Biophys. Acta, 1471: M43–M56, 2000.[Medline]
  70. Canman, C. E., Gilmer, T. M., Coutts, S. B., and Kastan, M. B. Growth factor modulation of p53-mediated growth arrest versus apoptosis. Genes Dev. 9: 600–611, 1995.[Abstract/Free Full Text]
  71. Waldman, T., Lengauer, C., Kinzler, K. W., and Vogelstein, B. Uncoupling of S phase and mitosis induced by anticancer agents in cells lacking p21. Nature, 381: 713–716, 1996.[Medline]
  72. Suzuki, A., Tsutomi, Y., Akahane, K., Araki, T., and Miura, M. Resistance to Fas-mediated apoptosis: activation of caspase 3 is regulated by cell cycle regulator p21WAF1 and IAP gene family ILP. Oncogene, 17: 931–939, 1998.[Medline]
  73. Shapiro, G. I., Koestner, D. A., Matranga, C. B., and Rollins, B. J. Flavopiridol induces cell cycle arrest and p53-independent apoptosis in non-small cell lung cancer cell lines. Clin. Cancer Res., 5: 2925–2938, 1999.[Abstract/Free Full Text]
  74. Lu, W., Lin, J., and Chen, J. Expression of p14ARF overcomes tumor resistance to p53. Cancer Res., 62: 1305–1310, 2002.[Abstract/Free Full Text]
  75. Raynal, S., Nocentini, S., Croisy, A., Lawrence, D. A., and Jullien, P. Transforming growth factor-ß1 enhances the lethal effects of DNA-damaging agents in a human lung-cancer cell line. Int. J. Cancer, 72: 356–361, 1997.[Medline]
  76. Lu, X., Errington, J., Curtin, N. J., Lunec, J., and Newell, D. R. The impact of p53 status on cellular sensitivity to antifolate drugs. Clin. Cancer Res., 7: 2114–2123, 2001.[Abstract/Free Full Text]
  77. Bronder, J. L. and Moran, R. G. Antifolates targeting purine synthesis allow entry of tumor cells into S phase regardless of p53 function. Cancer Res., 62: 5236–5241, 2002.[Abstract/Free Full Text]
  78. Krupenko, S. A., Wagner, C., and Cook, R. J. Expression, purification, and properties of the aldehyde dehydrogenase homologous carboxyl-terminal domain of rat 10-formyltetrahydrofolate dehydrogenase. J. Biol. Chem., 272: 10266–10272, 1997.[Abstract/Free Full Text]
  79. Osborne, C. B., Lowe, K. E., and Shane, B. Regulation of folate and one-carbon metabolism in mammalian cells. J. Biol. Chem. 268: 21657–21664, 1993.[Abstract/Free Full Text]
  80. Landberg, G. and Roos, G. Antibodies to proliferating nuclear antigen as S-phase probes in flow cytometric cell cycle analysis. Cancer Res., 51: 4570–4574, 1991.[Abstract/Free Full Text]



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