INTRODUCTION

Top Abstract Introduction Materials and Methods Results Discussion References

Topoisomerase (topo) II enzymes function to unknot and decatenate covalently closed circles of DNA. In mammalian cells a requirement for type II topo has been suggested in many aspects of DNA metabolism including replication and recombination (41). There are two isozymes of topo II with molecular masses of 170 and 180 kDa, termed topo II and topo II (7). While the biochemical activities of the two proteins are closely related, their cellular distribution and expression characteristics differ greatly. topo II is expressed at low levels in quiescent cells and is induced when cells are stimulated to enter the growth phase (32). As cells become contact inhibited, the levels of topo II are dramatically reduced (2, 22, 24). On the other hand, topo II expression does not correlate with proliferative status, remaining at constant levels in quiescent and proliferating cells (32, 39).

topo II is known to be expressed at high levels in tumors at both the protein and mRNA levels (19). This elevation is identified in a wide variety of tumors and is due in part to the increased growth fraction (3, 11, 17, 26). The expression of topo II is also increased as cells pass through the cell cycle (15), although reports vary as to the extent. In NIH 3T3 cells transformed by oncogenic ras (42) topo II levels increase, suggesting that oncogenic signaling might directly stimulate the topo II promoter, leading to increased protein levels in transformed cells. In support of this possibility, we found that oncogenic Ras is able to increase the expression of a reporter plasmid containing the basal topo II promoter driving a luciferase reporter gene (6). This increase, which requires ERK and JNK activities, was independent of cell cycle position (6).

Anti-topo II drugs are used clinically against a wide range of tumors (18) and target both isotypes of topo II, although a broad range of evidence indicates that topo II is the primary target (4). A number of studies have identified a close relationship between topo II levels and drug sensitivity (9, 12, 14, 17, 30, 31). Similarly, drug resistance is commonly correlated with decreased topo II levels (10, 28). On the other hand, factors other than topo II levels have been found to influence toxicity, including drug uptake, topo II phosphorylation (13, 16), and cellular factors involved in activation of cell death pathways (25, 28).

In order to better understand the interplay of cellular factors involved in controlling cellular response to drug treatment, we designed a series of experiments to analyze the roles of the cell cycle, oncogene action, and topo II levels independently of one another during treatment with an anti-topo II drug. While the level of topo II within the cell is believed to play a central role in determining drug toxicity (10, 36, 38, 40, 44), the considerations discussed above suggest that these levels might be altered by cell cycle position and oncogene activity. At the same time, these two physiological factors might directly influence drug toxicity independently of alterations in topo II levels (1, 23, 43). The challenge, therefore, is to separate the influence of the cell cycle and oncogene action on drug toxicity directly from their ability to alter topo II levels, and thereby to influence drug toxicity indirectly.

To address this question, we have developed a means to study the cell cycle expression characteristics of topo II in the presence and absence of oncogenic signaling. In addition, we have developed a means to determine cell cycle-related drug toxicity. The technical approach used relied on studies of continuously cycling cells. No attempt was made to synchronize cells in the cell cycle; rather, the cell cycle position of each individual cell was determined using either quantitative fluorescence analysis of DNA or time-lapse observations to determine the timing of mitoses (37). This technical approach has proven effective in previous studies of cyclin D1, where the expression characteristics of this protein were accurately determined in each cell cycle phase, together with the dependence of this expression on cellular Ras activity (20, 21). In the present study the same approach was utilized to determine the effect of the cell cycle and oncogene activity on the expression levels of topo II. We were somewhat surprised to learn that neither of these factors altered topo II protein expression levels in actively cycling cells. It was, therefore, possible to accurately determine the influence of cell cycle position and oncogene activity on the outcome of drug treatment without concern for topo II protein expression levels.

	MATERIALS AND METHODS

Top Abstract Introduction Materials and Methods Results Discussion References

Cell culture. All cells were cultured in Dulbecco modified Eagle medium supplemented with 10% calf serum or fetal calf serum and antibiotics. Cells were routinely cultured in a CO₂ incubator until the time of video analysis, at which time they were transferred to a moist chamber with a constantly refreshed, moistened 5% CO₂ atmosphere at 37°C. The cells proliferated as well in the environmental chamber used for video analysis as in the incubator for up to 8 days (at which time the cells had become completely confluent in each environment). Cells (5 × 10⁴ to 20 × 10⁴) were plated on a coverslip in a 35-mm-diameter dish in preparation for video analysis. For staining purposes cells were approximately 30% confluent, while for drug toxicity studies cells were approximately 10% confluent.

Time-lapse analysis. Time-lapse analysis was performed with a CCD camera attached to a frame capture board controlled by the National Institutes of Health Image program as previously described (37). The area of analysis was marked with two contiguous circles of different sizes using a diamond object marker (Leitz) prior to the beginning of the analysis (20). This allowed the viewing of the same area of the coverslip before and after addition of thymidine and allowed realignment of the area of analysis and identification of individual cells following staining and autoradiography. Oncogenic Ras (Leu61; 1 mg/ml) was microinjected into all the cells and only the cells within the designated circular area. There was no evidence of adverse effects of the injection or introduced protein in these analyses for more than 1 h, as judged by either morphological changes or alterations in the rate of cell division. As a control, nonspecific rat immunoglobulin (10 mg/ml) was injected into cells exactly as described above. The average cell cycle time of 16 to 18 h was increased by approximately 2 h during the period in which the cells were removed from the environmental chamber and microinjected. This lengthening of the cell cycle was due to a combination of the adverse effects of the microinjection and the reduction in the temperature of the culture during microinjection (which was performed at room temperature). In the first full cell cycle following microinjection the cells returned to the typical cell cycle time of unperturbed cultures.

Microinjection and antibody staining. The procedure for microinjection was identical to that previously reported (21). Cells to be injected were plated on a 22-mm-square coverslip and cultured in a 35-mm-diameter plate. The cells were inverted in culture medium during injection and placed on the stage of an upright microscope. The micropipette (outside diameter of less than 1 µm) was positioned within the culture medium just below the coverslip, and injections were performed by bringing the micropipette up into the cell while forcing the outflow of sample. Prior to injection the circles described above were scribed on the back side of the coverslip with a Leitz object marker, and the injections into all the cells enclosed by this circle were performed. Immediately following injection the coverslip was placed in the environmental chamber and time-lapse analysis was initiated. Injected cells were also identified by staining with an antibody against nonspecific rat immunoglobulin (10 mg/ml) included as a marker in Ras injections. The topo II levels were determined with a monoclonal antibody specific to the p170 alpha subunit of human topo II (catalog no. 2010-1; Topogen, Columbus, Ohio). The cells were counterstained with a Cy3-linked secondary antibody. Nuclei were isolated from NIH 3T3 cells (or these cells transformed by src, ras, or raf) following swelling in hypotonic buffer (10⁷ cells/ml; 5 mM KH₂PO₄ [pH 7.2], 2 mM MgCl₂, 10 mM -mercaptoethanol, 0.1 mM Na₂EDTA, protease inhibitors). The cells were broken with 20 strokes with a Dounce homogenizer on ice and collected by centrifugation at 1,000 × g for 6 min.

Analysis of data. Two forms of data presentation are utilized. The first consists of raw data in which the results for each cell are presented. To assess the results of numerous determinations, average values are also presented. To obtain average numbers, all the cells within a single analysis were considered to determine the G₁ and G₂ phase DNA readings. Each cell was then given a DNA reading based on this range for the specific experiment. Then, the topo II/DNA ratio for each cell was determined, and the average of all cells within the experiment was computed. The topo II/DNA ratio for each cell was then divided by the average for all cells in a given experiment. In this way, the DNA and topo II/DNA ratio readings for each cell were normalized for a given experiment. This made it possible to compare the results for different experiments even though the staining intensity in different experiments might vary slightly. For average determinations, the average for each experiment in a given DNA range was determined and then the values from all experiments were compared to determine the mean topo II/DNA ratio ± the standard error (SE) for all the experiments in a given DNA range. This calculation was performed for cells in all DNA ranges, and these numbers were then plotted versus the DNA level (Fig. 3A). Similar calculations were performed to compare the ratios of topo II levels in injected cells to those in uninjected cells (Fig. 3C) and the average DNA or topo II levels in time-lapse analyses (Fig. 4C). For the comparison of Western blotting and fluorescence readings the procedure was as previously described (20), except that the Western analysis was performed by blotting with an anti-topo II antibody followed by a horseradish peroxidase-coupled antimouse antibody coupled to ECL (Amersham). The luciferase levels were recorded on film and quantitated using the National Institutes of Health Image program. For comparison, average fluorescence readings were determined at each time period by staining with the fluorescent anti-topo II stain or by staining DNA with DAPI and quantitating the levels of fluorescence in the nuclear regions of over 250 cells. These readings were then averaged to obtain the values reported in Fig. 1.

	RESULTS

Top Abstract Introduction Materials and Methods Results Discussion References

Our goal is to determine the effects of the cell cycle, oncogenic activity, and topo II protein levels on the outcome of treatment with the anti-topo II drug etoposide. It was first necessary to determine exactly how topo II protein levels are affected by the cell cycle and oncogene action. This required a detailed cell cycle analysis of topo II protein levels. Our approach was to determine the cell cycle positions of individual cells in an asynchronous culture. This approach not only allowed analysis of cells in all cell cycle phases simultaneously but also eliminated any potential complication associated with methods of cell cycle synchronization. The cell cycle phase was determined in two separate ways. In the first, cells were fixed and DNA was stained with DAPI. Previous studies have conclusively demonstrated that the fluorescence intensity of the cell stained in this way is proportional to DNA content. Moreover, the analysis of monolayer cells was found to be extremely accurate due to the high-resolution optics involved, the ability to analyze fluorescence specifically within the nuclear region, and the low level of cytoplasmic interference within the nuclear region in flattened monolayer cells (20). In the second technique, cell cycle position was assessed by monitoring cells in time-lapse for 24 h and then determining the age of each cell (or the time since passing through mitosis) at the time of fixation (21).

To determine the level of topo II protein in each cell, the cells were stained with an antibody specific to topo II. It was first necessary to demonstrate that this staining procedure resulted in fluorescence levels proportional to those of topo II protein within the cell. To accomplish this, topo II protein levels and fluorescence were compared in cells synchronized by serum deprivation and restimulation. At various times following serum addition to quiescent NIH 3T3 cultures, lysates were collected for Western analysis, while cells in parallel cultures were fixed and stained for topo II and DNA. The amount of topo II protein at each time point was determined by quantitating the intensity of the Western band. This was then compared to fluorescence intensity by measuring the fluorescence of 200 to 400 cells in a parallel culture and determining the average fluorescence (21). The results of a typical analysis (Fig. 1) indicate a close correlation between the average fluorescence intensity of the topo II stain and the absolute amount of topo II protein within the culture. In separate analyses, it was found that only full-length topo II was present in isolated nuclei, thus reducing the possibility that the results obtained reflected partially degraded protein (data not shown). It is, therefore, possible to measure fluorescence intensity and determine the relative amounts of topo II in individual cells.

View larger version (19K): [in this window] [in a new window]   FIG. 1.   Comparison of topo II protein levels and fluorescence intensity. NIH 3T3 cells were rendered quiescent by culture in low serum (0.5% calf serum) for 48 h. Serum was then added to separate cultures, and at the times indicated lysates were prepared for quantitative Western analysis. For comparison, a parallel plate was fixed and stained for topo II with a specific, indirect fluorescent stain. Images of the stained cultures were collected with a CCD camera (together with images of the DNA stained with DAPI). The total amount of topo II-associated fluorescence was determined for each nucleus. The average intensity of 200 to 400 individual nuclei is plotted versus the relative amount of topo II protein determined by quantitating the topo II protein band in Western analysis.

Increase of topo II protein through the cell cycle. The levels of topo II through the cell cycle of actively cycling cells were next determined by fixing and staining an asynchronous culture of NIH 3T3 cells for topo II and DNA as described above. Separate images of each fluorochrome were collected from the same cells. The nuclear region of each cell was identified from the DNA stain, after which the total amounts of DNA- and topo II-associated fluorescence for each nucleus were determined. When the DNA fluorescence levels for individual cells are plotted versus topo II-associated fluorescence, it can be seen that the amounts of topo II continuously increased as the cells progress from G₁ to G₂ phase (Fig. 2A). While the amounts of topo II protein increase during passage through the cell cycle, it was somewhat surprising, based on previously reported data, that the increase was not larger than the one observed. Most cell constituents passively increase in amount during the cell cycle due merely to the increase in cell size and mass. It is critical to determine if this might be the explanation for the slight increase in topo II protein observed above.

View larger version (36K): [in this window] [in a new window]   FIG. 2.   Topo II expression through the cell cycle. (A) An asynchronous culture of NIH 3T3 cells was fixed and stained for DNA and topo II. The levels of each fluorochrome were determined by image analysis, and the DNA levels for each cell were plotted versus the topo II levels, with the G1 and G2 levels of DNA indicated. (B) To determine relative topo II levels, the topo II-associated fluorescence value was divided by the DNA-associated fluorescence and this ratio was plotted versus the DNA level (and therefore cell cycle position).

View larger version (23K): [in this window] [in a new window]   FIG. 3.   Topo II levels through the cell cycle. (A) The topo II and DNA fluorescence levels of individual cells were determined in nine separate experiments similar to that shown in Fig. 2B. The ratio of topo II level to DNA level for each cell was determined, and the ratio of this value to the average value for all the cells in a given experiment is reported. This number, therefore, indicates the relative topo II/DNA ratio for each cell compared to those for all the cells of a given experiment. A cell with a value of 1.0 would have an average topo II/DNA ratio in that experiment. For comparison, cells of each experiment were divided into groups according to their DNA content, and the average topo II/DNA ratio for each group was determined. Groups with the same DNA content from all nine experiments were then compared to determine the mean ratio of topo II level to DNA level (± SE) for this DNA range. This value was then plotted versus the DNA level. (B) Oncogenic Ras (Leu61; 1 mg/ml) was microinjected into cells within an asynchronous NIH 3T3 culture which was incubated for 24 h, fixed, and stained for topo II and DNA. DNA-associated fluorescence is plotted versus topo II-associated fluorescence. (C) Results from six separate experiments like those shown in panel B were compared. Cells in each experiment were divided into groups according to their DNA content, and the average topo II level for injected cells in a group was divided by the average level for uninjected cells. The ratios obtained in this way for all experiments were then combined, and the mean ratio of the topo II level in injected cells to that in uninjected cells (± SE) for each DNA range was determined. This value was plotted versus the DNA content for each group. Because there was little difference between cells injected with oncogenic Ras and uninjected cells, data concerning control injections are not shown.

Time-lapse analysis of topo II levels. To confirm the above results, topo II expression through the cell cycle was analyzed using time-lapse to determine cell cycle position. Cells of an asynchronous culture were monitored for 25 h prior to fixation and staining for topo II. The level of topo II-associated fluorescence was determined for each cell as described above. Individual cells were then monitored in the time-lapse movie to determine their ages, or how long prior to fixation they had passed through mitosis (29). In previous studies we have shown that most NIH 3T3 cells less than 5 h old are in G₁ phase, that cells 5 to 12 h old are in S phase, that cells older than 12 h are most likely in G₂ phase, and that the average generation time was 16 to 17 h (20, 37). As an indication of the cell cycle characteristics of these cultures, the ages of individual cells in a typical analysis are plotted versus their DNA contents (Fig. 4A). Although most cells in the culture behave as described above, a small number display slower transit through the cell cycle. The topo II-associated fluorescence also increased with age (Fig. 4B). To relate topo II levels to DNA levels as these cells progressed through the cell cycle, cells were grouped according to age and the average topo II fluorescence levels for each group were determined, as was the average DNA fluorescence level. When the average topo II and the average DNA levels were plotted versus age, it was clear that topo II levels and DNA content increased together through the cell cycle in this time-lapse analysis (Fig. 4C). Based on all the above data, therefore, we conclude that the topo II content of cycling cells increases roughly in proportion to the increase in DNA content through the cell cycle in all cell types analyzed.

View larger version (39K): [in this window] [in a new window]   FIG. 4.   Time-lapse studies of topo II through the cell cycle. Cells were monitored in time-lapse for 25 h, fixed, and stained for topo II and DNA. These levels for each individual cell were then determined by photography and image analysis as described previously. The age of each cell was determined by analysis of the time-lapse movie. (A) The age of each cell plotted versus its DNA content. (B) The topo II content of each cell plotted versus its age. (C) Cells were grouped according to age, and the average DNA and topo II contents were determined and plotted versus age. It is clear that while most cells cycle as expected, a small population progress through the cell cycle much more slowly. The cells which failed to divide during the entire time-lapse analysis are displayed as having an age of 25 h.

Ras injection and topo II levels. We next analyzed the effects of constitutive oncogenic signaling on topo II levels. Oncogenic Ras (L61; 1 mg/ml) was microinjected into cells of an asynchronous culture, and the topo II and DNA levels were determined 24 h later by fluorescence photography and image analysis. No increase in the topo II levels was observed following Ras injection during any cell cycle phase (Fig. 3B). As above, for a critical comparison the results of several separate experiments were combined. Cells in individual experiments were divided into groups according to DNA content. For each group the topo II levels for all Ras-injected cells were compared to topo II levels for uninjected, neighboring cells. Results from all six experiments were combined to determine the mean ratio of the topo II level in injected cells to that in uninjected cells (± SE) for each DNA group (Fig. 3C). The topo II levels for injected and uninjected cells were remarkably similar, particularly during the first half of the cell cycle, although a slight reduction might have been induced by Ras during G₂ phase. Similar results were obtained when Ras-injected cells were compared to cells injected with rat immunoglobulin as a control.

Ras stimulation of topo II in slowly cycling cells. To confirm the above results, oncogenic Ras was injected into cells, which were then monitored for 25 h in time-lapse prior to fixation and analysis of topo II levels. As above, no overall stimulation of topo II levels by Ras could be observed (Fig. 4B). However, this analytical approach allowed the identification of cells with lengthened cell cycle phases. Cells which failed to divide in the 25 h of the time-lapse observation are represented as having an age of 25 h. The topo II levels in these cells were low as expected, except in the Ras-injected cells. In some of these slowly cycling, Ras-injected cells the topo II levels increased dramatically (Fig. 4B). Such cells would not have been apparent without time-lapse analysis. These cells are considered potentially interesting because they might serve as a culture model for tumor cells. Like most cells of a tumor, they retain proliferative capacity but cycle slowly. The fact that these cells responded to Ras injection by increasing topo II levels, therefore, is an observation which was carefully verified.

View larger version (38K): [in this window] [in a new window]   FIG. 5.   Ras stimulation of topo II in slowly cycling NIH 3T3 cells. (A) NIH 3T3 cells and a clone of these transformed by oncogenic ras (Val12) were labeled with [3H]thymidine for 20 h, fixed, stained for topo II, and autoradiographed. The topo II levels of individual slowly cycling cells, which failed to incorporate thymidine, in both ras-transformed and untransformed cultures were determined, and the means (± SE) are presented. For comparison, the average topo II level of 200 cycling, thymidine-labeled cells in each culture was also determined. (B) src-transformed NIH 3T3 cells were labeled with BrdU for 24 h, fixed, and stained for BrdU and topo II. The topo II-associated fluorescence is plotted versus the BrdU-associated fluorescence for all cells in the culture. Slowly cycling cells form a line along the bottom of the profile with low BrdU staining.

Drug toxicity and the cell cycle. Once the effect of the cell cycle and oncogenic activity on topo II levels was determined, it was possible to determine the influence of each of these factors on drug toxicity. This comparison was simplified by the fact that neither the cell cycle nor oncogenic activity altered topo II levels, so that the effect of each could be determined without concern that secondary effects on topo II levels were involved. Drug toxicity studies were performed in asynchronous cultures with two consecutive time-lapse movies of the same cells separated by a brief (80-min) treatment with etoposide (2 to 10 µM). From the first movie (24 h) the cell cycle position of each individual cell at the time of drug treatment was determined, while from the second movie (50 to 75 h) the fate of each treated cell was analyzed (Fig. 6). This allowed analysis of the effects of the cell cycle on drug toxicity without treatments required to induce cell cycle synchrony. Cells were poisoned with 5-azacytidine (100 to 500 µM) or tritiated thymidine (2 mCi/ml) (8, 34) as controls in separate analyses.

View larger version (15K): [in this window] [in a new window]   FIG. 6.   Experimental procedure for cell cycle toxicity analysis. In order to determine the effects of the cell cycle on drug toxicity, NIH 3T3 cells were monitored in time-lapse for 25 h, the final 80 min of which was in the presence of the drug. The drug was removed from the culture, and a second movie of 50 to 70 h was immediately initiated, making certain that the same cells were monitored throughout. Each cell is illustrated by a single horizontal line, with the branch point representing mitosis. Age is the time between the initiation of drug treatment and the preceding mitosis. The fates of treated cells are determined during the second movie, with the most common fates illustrated.

View larger version (113K): [in this window] [in a new window]   FIG. 7.   Rapid cell death and abortive mitosis. (Top) A small proportion of NIH 3T3 cells treated with etoposide experienced a sudden, dramatic cell death. A typical event is pictured, with the relative times indicated. Note that the most dramatic effects took place between frames 8 min apart. (Bottom) A similar presentation of a cell passing through an abortive mitosis following etoposide treatment is made, with the relative times indicated. (A video presentation of each of these events is presented at http://www.lerner.ccf.org/labs/stacey/dir.cgi, labeled RD2 and AM1).

View larger version (15K): [in this window] [in a new window]   FIG. 8.   Cell cycle and drug toxicity. (A) Six individual experiments were performed as described for Fig. 6. In these experiments NIH 3T3 cells were treated with etoposide (7 µM) to determine cell cycle-specific toxicity. The age and fate of all treated cells were determined, the cells were separated according to age, and the proportion of normal cells for each age period was calculated. The results for all six separate experiments for each age range were then compared to yield the mean percentage of normal cells (± SE) for cells of a given age. This number is plotted versus the age at the time of treatment. (B) A similar analysis was tabulated from five separate experiments using src-transformed NIH 3T3 cells treated with 2 µM etoposide. (C) NIH 3T3 cells were treated with 5-azacytidine (150 µM) in three separate experiments. (D) Two separate experiments in which NIH 3T3 cells were treated for 80 min with [3H]thymidine (2.5 µCi/ml; 79 Ci/mM) were performed. (E) The results from panels A and D are plotted together for comparison.

View larger version (29K): [in this window] [in a new window]   FIG. 9.   Drug toxicity after mitotic selection. At 90-min intervals mitotic NIH 3T3 cells were detached mechanically from asynchronous cultures and replated in several parallel plates. At the indicated times following replating each culture was treated with the indicated concentration of etoposide (A) or 5-azacytidine (AC) (B) for 80 min. The cultures were washed and cultured for an additional 3 days before trypsinization and counting. The numbers reported indicate the numbers of cells compared to those for mock-treated cultures. (C) To determine cell cycle characteristics, cells were labeled with tritiated thymidine at various times following replating, immediately fixed, and autoradiographed to determine the proportion of cells in S phase.

Oncogenic signaling and toxicity. Experiments were last performed to determine the effect of oncogenic activity on etoposide toxicity. Experiments similar to those described above were repeated with NIH 3T3 cells transformed by the src oncogene. As previously reported (5), these cells were much more sensitive to etoposide toxicity than untransformed cells. Experiments were performed with 2 µM etoposide so that the proportion of cells killed would be approximately equal to that observed when untransformed cells were treated with 7 µM etoposide. While the sensitivity of src-transformed cells increased, the overall cell cycle characteristics of killing were similar to those observed with untransformed cells, with the possible exception that the overall cell cycle effects of toxicity might have been reduced in the transformed cells (Fig. 8B). To extend this observation, NIH 3T3 cells were injected with oncogenic Ras protein in a defined area of the culture and the toxicity of etoposide (2 µM) was determined with the time-lapse analysis. In this case the overall proportions of normal cells for the injected and uninjected neighboring cells were determined. As a control, similar experiments were performed by microinjecting rat immunoglobulin as a negative control. In repeated experiments, a dramatically increased sensitivity of Ras-containing cells was observed. The proportion of poisoned cells increased over threefold following Ras injection (Fig. 10A). Unfortunately, the overall numbers of these cells were too small to make definite conclusions regarding the cell cycle profile of toxicity following Ras injection. Since we know that the overall levels of topo II do not increase following Ras injection, this clearly indicates that oncogenic activity sensitizes the cells to etoposide independently of alterations in topo II levels.

View larger version (21K): [in this window] [in a new window]   FIG. 10.   Oncogenic transformation and etoposide toxicity. (A) The time-lapse analysis of toxicity (Fig. 6) was applied to cells injected with oncogenic Ras (Leu61; 1 mg/ml). The proportion of the cells which divided normally or which were killed was determined in three separate experiments, and the combined results are presented. For comparison, the results with uninjected, neighboring cells are also presented, together with the results following microinjection of a control protein (10 mg of rat immunoglobulin/ml). (B) Survival of slowly cycling cells following drug treatment. Cells which failed to divide in the movie prior to drug treatment from all experiments presented in Fig. 8A to C are considered. The proportion of these cells which divided normally is presented, together with the proportion of cells in S phase at the time of drug treatment which remained normal following treatment. Results for NIH 3T3 cells treated with 7 µM etoposide, src-transformed NIH 3T3 cells treated with 2 µM etoposide, and NIH 3T3 cells treated with 150 µM 5-azacyticine are presented. These results are based on a total of 67 nondividing cells in all experiments.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

A number of technical approaches were developed to separately assess the roles of the cell cycle and oncogenic signaling in the sensitivity to the anti-topo II drug etoposide, independently of changes in topo II levels. It was first necessary to determine the influence of the cell cycle and oncogene activity on topo II expression. In these studies we avoided working with synchronized cell populations to avoid potential complications resulting from treatments required to induce cell cycle synchrony. Instead, an analytical approach was utilized to accurately assess cell cycle-specific protein expression levels in asynchronous cultures by determining the cell cycle positions of individual cells using quantitative fluorescence staining of DNA or time-lapse analysis. This approach has proven useful in analyzing the cell cycle expression levels of cyclin D1 protein and in determining the signaling requirements for this cell cycle-regulated expression pattern (21). In the present study, it was first necessary to directly demonstrate that the fluorescence intensity of cells stained with a specific antibody stain against topo II accurately reflected the amount of protein within the cell. Once this relationship was established, it was possible to determine topo II expression characteristics of individual cells. We were surprised to find that there was no specific increase in topo II protein as cells progress through the cell cycle. This was the case for NIH 3T3 cells as well as for all other cell lines analyzed (including HeLa cells, T24 [bladder tumor] cells, MCF-7 and 468 [breast tumor lines] cells, and MRC-5 diploid human fibroblasts). It is not clear if topo II levels are linked directly to DNA levels within the cell or if another related cellular factor is involved. Moreover, contrary to our initial beliefs, there was absolutely no induction of topo II levels in oncogene-containing cells. These observations conflict with some published reports most probably because we have studied only actively cycling cells. In studies with serum-deprived cells, the addition of serum induces not only a change in the cell cycle but also, more importantly, a total change in growth phase. Topo II is known to be tremendously sensitive to changes in growth state, with high levels of expression in proliferating cells (2, 43).

The only exception to this pattern was observed in cells within the asynchronous culture which exhibited an extended cell cycle time. In perhaps 20% of these cells oncogenic Ras and src were able to induce high levels of topo II protein expression. To explain these results, we predict that topo II levels are strictly tied to the level of a cellular factor which increases proportionally through the cell cycle. Our studies have conclusively shown, however, that a portion of the topo II promoter is responsive to Ras activity (6). In cells which remain viable but which are temporarily removed from active cycling this portion of the topo II promoter apparently becomes dominant and the cell responds by producing high levels of topo II protein. This fact might be responsible for the high levels of topo II protein within some tumors (10) which, like the slowly cycling NIH 3T3 cells, retain proliferative capacity but which are temporarily not passing actively through the cell cycle. Indeed, the slowly cycling cells in this study, like many tumor cells, were able to reenter the pool of actively cycling cells at any time, as indicated by the fact that (except in transformed cells) they were as likely as the general cellular population to divide normally following drug treatment.

The fact that neither the cell cycle nor oncogene action influenced topo II levels in cycling NIH 3T3 made it possible to study the effects of these two factors on drug toxicity, without concern for alterations in topo II levels. Previous work with synchronized cells pointed to a close correlation between S phase and drug toxicity (30). In these studies, drug treatment between two time-lapse observations allowed determination of the cell cycle phase at the time of drug treatment and the effects of this treatment on individual cells. The results of numerous studies established that etoposide was approximately threefold more toxic when added to cells in late S and G₂ phases than when added in G₁ phase. (Similar results were obtained with cells collected by mitotic detachment.) The exact cell cycle timing of this sensitive period was determined and compared to that for treatment with labeled thymidine. The highly selective killing of cells only in S phase following tritiated thymidine treatment was remarkable and emphasizes the validity of our procedure for determining cell cycle-related toxicity. Finally, the cell cycle toxicity pattern was specific to etoposide, as it was not seen with another toxic agent, 5-azacytidine. While a 10% increase in the levels of topo II through the cell cycle exactly correlated with the period of increased drug sensitivity (Fig. 3A), we are skeptical that this small difference could account for the dramatic differences in toxicity observed (Fig. 8A). Moreover, the alterations in drug sensitivity described below with oncogene-containing cells could not be accounted for by even slight alterations in topo II levels. We have not considered the potential role of topo II in these studies, because the expression of this protein is not increased as cells enter a growing phase (35), even though there is evidence that it can serve as a target of anti-topo II drugs (10, 33).

While further study is required, we consider it significant that the topo II expression characteristics of slowly cycling cells were so different from those seen in actively proliferating cells. An effective antitumor treatment must target cells which are not actively cycling, since these make up the majority of all tumors. This study demonstrates that such cells, which are neither actively cycling nor quiescent, are likely to have unique biological characteristics. A study of such cells might be highly informative relative to the behavior of cells within a tumor.

When src-transformed cells were analyzed, it was clear, as previously reported, that oncogenic transformation increased the sensitivity to etoposide. Approximately equal killing was obtained in transformed cells with etoposide at 30% of the level used with untransformed cells. Interestingly, while the cell cycle effects of toxicity might have been diminished somewhat in transformed cells, transformed cells showed the same general pattern of increased toxicity late in the cell cycle as untransformed cells. Moreover, increased toxicity was observed in the slowly cycling cells transformed by oncogenic src, despite the fact that only a small number of these cells had increased topo II protein levels. To eliminate the possibility that clonal effects might have been responsible for the increased sensitivity with src-transformed NIH 3T3 cells, microinjection of oncogenic Ras was also shown to dramatically increase toxicity to etoposide. It is, therefore, concluded that the action of oncogenes from two separate classes is able to dramatically sensitize NIH 3T3 cells to etoposide toxicity, even though no changes in topo II levels result. This result is similar to that of a recent study of tumor cells where the presence of an oncogenic ras mutation correlated to sensitivity to anti-topo II drugs (27).

We conclude, therefore, that both the cell cycle and oncogene action play significant roles in determination of the effect of treatment with etoposide, without altering topo II expression levels. The explanation for this observation is not presently known. It is possible that these two physiological factors might influence the phosphorylation of topo II or its intermolecular associations. On the other hand, cell cycle and oncogene action might affect a molecule totally distinct from topo II, which plays a significant role in controlling drug toxicity. Further study will be required to answer these questions.