Programmed cell death and mutation induction in AHH-1 human lymphoblastoid cells exposed to m-amsa

Fundamental and Molecular Mechanisms of Mutagenesis

ELSEVIER

Programmed

Mutation Research 329 (199.5) 79-96

cell death and mutation induction in AHH-1 human lymphoblastoid cells exposed to m-amsa

Suzanne M. Morris a!*, Olen E. Domon a, Lynda J. McGarrity Daniel A. Casciano a

a, James J. Chen b,

a Divison of Genetic Toxicology, Department of Health and Human Services, US Public Health Service, Food and Drug Administration, National Center for Toxicological Research, 3900 NCTR Road, Jefferson, AR 72079, USA b Division of Biometry and Risk Assessment, Department of Health and Human Services, US Public Health Service, Food and Drug Administration, National Center for Toxicological Research, 3900 NCTR Road, Jefferson, AR 72079, USA

Received 16 December 1994; accepted 15 February 1995

Abstract One role of programmed cell death (apoptosis) is the removal of cells with DNA damage from the population. Certain cells, however, are able to suppress the signals for apoptotic cell death and maintain viability. This suggests that the susceptibility of a cell to either undergo apoptosis or escape from the apoptotic death pathways may be an important factor in chemical mutagenesis. In order to provide insight into the role of apoptosis in the recovery of chemically induced mutants, AHH-1 cells were exposed to the chromosomal mutagen, m-amsa, and the percentage of cells undergoing apoptosis or necrosis quantified by flow cytometry. Logistic regression analysis revealed that the primary manner of cell death was by apoptosis. Two specific-locus mutations assays, the tk and the hprt, were utilized as markers for cells with DNA damage and that retained clonogenicity under conditions known to induce apoptosis. Analysis of variance indicated that the concentration-dependent increase in the mutant fraction at the tk locus was significant and the result of the recovery of clones with the slow-growth phenotype. Because this phenotype is thought to reflect chromosomal mutations, these results are consistent with the survival and clonogenicity of damaged cells. This suggests that the ability to recover mutant cells may be influenced by the suppression of or an escape from the apoptotic death pathways. Keywords:

Lymphoblastoid

cells; Human;

Apoptosis;

tk mutation;

Abbreviations: ANOVA, analysis of variance; EMS, ethyl methanesulfonate; ENU, N-ethyl-N’-nitrosourea; FDA, fluorescein diacetate; FSC, forward scatter; hprt, hypoxanthine phosphoribosyl transferase; m-amsa, 4’-(9-acridinylamino)methanesulfon-m-aniside; PI, propidium iodide; SSC, side scatter; 6-TG, 6-thioguanine; tk, thymidine kinase; TFT, trifluorothymidine; pos, positive; neg, negative; I, resistant

hprt mutation;

m-Amsa

* Corresponding author. Tel. (501) 543-7580; Fax (501) 5437136.

0027-5107/95/$09.50 0 1995 Elsevier Science B.V. All rights reserved SSDI 0027-5107(95)00020-S

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1. Introduction

Apoptosis (programmed cell death) is a pathway for cell death that is morphologically and biochemically distinct from cellular necrosis. Classically, apoptosis is recognized as an effective mechanism by which cell numbers are regulated during development and differentiation. Recently, however, has come an understanding of the importance of programmed cell death in both the response to toxic insult and the progression of cells towards malignancy. Evidence indicates that pivotal roles exist for both oncogenes and tumor suppressor genes in the decision as to whether a cell proliferates or is recruited into the pathways for cell death (reviewed in Dive and Wyllie, 1993). The overexpression of an oncogene confers a selective advantage on the mutant cells in contrast to the advantage gained by the loss of function in a mutant tumor suppressor gene (reviewed in Oren, 1992). The most extensively studied relationship between oncogene overexpression and apoptosis is that of bcl-2 expression in B-cell lymphoid malignancies. In studies of cells derived from follicular lymphoma (Tsujimoto et al., 1985; Hockenberry et al., 1990), in cell lines transfected with the bcl-2 gene (Vaux et al., 1988; Tsujimoto, 1989a,b; Miyashita and Reed, 19921, and in mice transgenic for the bcl-Zig minigene (McDonnell et al., 1989, 19901, overexpression of the gene resulted in the inhibition of apoptosis in cells recruited into the bcl-Zdependent pathways (reviewed in Hickman, 1992; Korsemeyer, 1992a,b; Oren, 1992; McDonnell, 1993). The tumor suppressor gene, ~53, functions in the recruitment of damaged cells into the p53-dependent apoptotic cell death pathway (Yonish-Rouach et al., 1991), in addition to its role in the control of cell proliferation (reviewed in Donehower and Bradley, 1993). Resistance to death through apoptosis is a component of the mutant ~53 phenotype and thus, mutant cells are able to survive and proliferate (Lowe et al., 1993; Clarke et al., 1993). The recruitment of cells into these pathways provides an effective means of destroying damaged cells. Thus, a mutation in an oncogene or tumor supressor gene compromises the elimination of these cells from the population and results in the

Research 329 (I 995) 79-96

survival and proliferation rather than the death of damaged cells (Korsemeyer, 1992a,b; Lane, 1993). In a previous study (Morris et al., 1994a), exposure of AHH-1 human lymphoblastoid cells to the simple ethylating agents, ethyl methanesulfonate (EMS) and ethyl nitrosourea (ENU), resulted in the recruitment of cells into the apoptotic death pathways. In a subsequent study (Morris et al., 1994b), clones, isolated by resistance to 6-thioguanine (6-TG’) and presumed hypoxanthine phosphoribosyl transferase (hpti) mutants, were recovered at concentrations of EMS and ENU that induced apoptosis. Thus, cells that retained either DNA damage or the effects of that damage were able to suppress the signals for movement into the apoptotic cell death pathways. This was in contrast to the known role of apoptosis in the elimination of damaged cells from the population. The interpretation of the data from these experiments was limited by the fact that only the recovery of hprt mutations was studied and that this may not reflect the entire spectrum of DNA damage induced by exposure to EMS or ENU. Molecular analysis of the mutation spectrum of hprt mutants has revealed that mutation induction by EMS and ENU arises primarily from the formation of 02-ethylthymine, 04-ethylthymine and 06-ethylguanine (Bronstein et al., 1991; Li et al., 1992). However, DNA ethylation occurs at a variety of sites on chromosomal DNA and a multiplicity of ethylated adducts are formed (Beranek et al., 1980; Heflich et al., 1982). Thus, one explanation of our previous results is that while the mutagenicity of EMS and ENU resulted from oxygen alkylation, the signal for apoptotic cell death related to nitrogen alkylation and the formation of DNA adducts such as N’-ethylguanine or N3-ethyladenine. In order to extend these observations and to provide insight into the nature of the DNA damage present in cells that either escape from or suppress the signals for apoptosis, the thymidine kinase (tk) mutagenesis assay was utilized as a marker for cells with DNA damage in addition to the hprt assay. In mouse lymphoma cells (L5178Y) and human lymphoblastoid cells (TK61, chromo-

SM. Morris et al. /Mutation Research 329 (1995) 79-96

somal mutations can be phenotypically distinguished from point mutations on the basis of growth characteristics (Moore et al., 1985a; Clive, 1987; Liber et al., 1989). Presumed chromosomal mutations are recognized by the small colony (L5178Y) or slow growth phenotype (TK6), in contrast to the base-pair substitution mutations, frame-shift mutations and intragenic deletions associated with the large colony (L5178Y) or normal growth phenotype (TK6). Molecular (Applegate et al., 1990; Li et al., 1992) and cytological analyses (Hozier et al., 1981; Moore et al., 1985b) have revealed the chromosomal nature, primarily multi-locus deletions, of the DNA damage detected in mutants identified by the altered growth phenotype. In these experiments, the potent chemotherapeutic agent, 4’-(9-acridinylaminolmethanesulfonm-aniside (m-amsa), was selected for the following reasons. The mode of interaction with chromosomal DNA is well established; m-amsa intercalates into the DNA, preferentially at the A-T sequences, and stabilizes the cleavable complex. The cleavable complex forms when the individual homodimers of the enzyme, topoisomerase II (topo-III, bind to the cleaved DNA strands as DNA undergoes decatenation during synthesis (reviewed in Marshall and Ralph, 1985; Liu, 1989). Further, the efficient induction of tk mutants with the small coIony phenotype and the induction of chromosomal deletions in L5178Y cells exposed to m-amsa is consistent with the induction of chromosomal mutations by this compound (DeMarini et al., 1987; Doerr et al., 1989). The induction of multilocus deletions by m-amsa has recently been confirmed in a Chinese hamster ovary-human hybrid cell line by the recovery of Sl- mutants with deletions 1.5-2.0 mbp in length (Shibuya et al., 1994). In addition, recent evidence indicates that exposure to m-amsa results in the movement of cells into the pathways for programmed cell death (Del Bino and Darzynkiewicz, 1991; Del Bino et al., 1991; Walker et al., 1991; Gorczyca et al., 1993a,b,c). The ordered fragmentation of chromosomal DNA, consistent with cell death by apoptosis, was detected in m-amsa (and other topo-I and topo-II inhibitors&exposed cells of

81

lymphoid origin (Jaxel et al., 1988). That the DNA fragmentation was the result of the apoptosis-associated endonucleolysis and not due to a direct effect of cleavable complex stabilization was confirmed in the experiments of Schneider et al. (1989). The protein-linked single- and doublestrand breaks associated with cleavable complex stabilization were detected, but the DNA fragmentation and subsequent death did not occur in the presence of RNA and protein synthesis inhibitors known to block apoptotic cell death. The initial detection of 300 kb fragments, the subsequent appearance of 50 kb fragments and then, the detection of fragments that are multiples of 180 bp and form the traditional DNA ladder by pulsed-field and agarose gel electrophoresis were consistent with death through apoptosis (Walker et al., 1991, 1993; Oberhammer et al., 1993). The involvement of oncogenes in the signal transduction pathways leading to apoptotic cell death is not limited to bcl-2 and ~53. Expression of the oncogene, c-myc, occurs in cycling cells and sensitizes or ‘primes’ cells to either proliferate or undergo apoptosis (Evan et al., 1992; Dive and Wyllie, 1993). In RAT-l fibroblasts which constitutively overexpress c-myc, growth factor withdrawal by serum deprivation rapidly induced apoptosis and the sensitivity to undergo serum deprivation-induced apoptosis was correlated to the level of c-myc expression (Evan et al., 1992). In addition, recent studies in an interleukin-3 (IL-3)-dependent mast cell line indicate that the v-abl oncogene product is able to suppress apoptosis (Evans et al., 1993). Thus, the recovery of mutant clones may be affected not only by the loss of viability that results from chemical damage, but also by the inherent susceptibility of a specific genotype to undergo apoptosis. These findings led to our interest in further examining the role of programmed cell death in the mutagenicity of m-amsa in AHH-1 human lymphoblastoid cells. AHH-1 cells, a B-cell line transformed by Epstein-Barr virus, are heterozygous at the tk locus. Thus, AHH-1 cells were exposed to increasing concentrations of m-amsa and the resulting mutant fraction at the tk locus (both normal growth and slow growth phenotypes) and the hprt locus quantified. In order to

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determine if m-amsa exposure resulted in the recruitment of AHH-1 cells into the pathways for cell death, the percentages of apoptotic, necrotic and viable cells were determined flow cytometritally with the propidium iodide (PI)/fluorescein diacetate (FDA) assay (Pavlik et al., 1984; Ross et al., 1989). Cells undergoing apoptosis will be PI negFDAneg, late apoptotic and necrotic cells will be PIPoSFDAneg, and viable cells will be PInegFDAPoS. These classifications of AHH-1 cells have been confirmed by electronically sorting the populations and morphological examination of the hematoxylin/eosin-stained cells (Morris et al., 1994a) and electron microscopy of osmium tetroxide-fixed cells (Morris, SM., L.J. McGarrity, O.E. Domon et al., in preparation). Exposure to m-amsa was conducted in both serum-free medium and complete medium as a means of determining the effect of serum deprivation on mutant recovery and cell death in AHH-1 lymphoblastoid cells.

in complete medium for a minimum before chemical exposure.

of 72 h

Chemicaki and exposure conditions

m-Amsa was the generous gift of Dr. David DeMarini, USEPA, Research Triangle Park+,NC. In order to expose cells to m-amsa, the cells were pelleted by gentle centrifugation, the supernatant removed and the cells enumerated by hemacytometer. Cells were reseeded at a density of 5 X lo5 cells per ml in 60 ml of medium RPM1 1640 (Gibco) supplemented with 1% L-glutamine (Gibco) and 1% penicillin-streptomycin (Gibco). m-Amsa was diluted in DMSO (Sigma, St. Louis) and each culture (including the control) received 300 ~1 of the diluent. The 24-h exposure of cells to m-amsa was conducted both with 10% serum and without serum. The chemical was removed by gentle centrifugation and the cells were enumerated by hemacytometer and seeded at a density of 5 x lo5 cells per ml in complete medium. Cultures were maintained at 37” C in a 5% CO,, 100% humidified atmosphere.

2. Materials and methods Mutation assays Cell culture

AHH-1 cells, a human lymphoblastoid cell line, were obtained under license from Gentest Corporation (Woburn, MA). The cells were routinely cultured in medium RPM1 1640 (Gibco, Grand Island, NY) supplemented with 10% iron-supplemented calf serum (Hyclone, Logan, UT), 1% L-glutamine (Gibco), and 1% penicillin-streptomycin (Gibco), at 37°C in a 5% CO,, 100% humidified atmosphere. Cells were maintained at a density of 2-5 x lo5 cells/ml and the medium was replenished at 48-72-h intervals. In order to reduce the spontaneous mutation frequency, cells were cultured in antibiotic-free medium RPM1 1640 containing 10% iron-supplemented calf serum, 1% r_-glutamine and 2 X low4 M hypoxanthine, 8 X lo-’ M aminopterin and 3.2 X lop5 M thymidine (HAT supplement, Gibco) for 72 h. After removal from HAT medium, the cells were cultured in antibiotic-free medium RPM1 1640 with 10% ISCS, 1% L-glutamine and 2 X 10V4 M hypoxanthine and 3.2 X low5 M thymidine (HT supplement, Gibco) for 24 h. Cells were cultured

Determination

of the mutant fraction at the

hprt locus was as described in Penman and Crespi (1987) and at the tk locus as described in Crespi

et al. (1985) with the modifications suggested by Liber et al. (1989) for the discrimination of slow growth mutants in TK6 cells. Briefly, 5 x lo5 cells per ml were exposed in either complete medium or medium without serum to m-amsa at 0.0, 1.0, 10.0, 50.0, 100.0 or 250.0 ng/ml for 24 h. After exposure, cells were cultured in complete medium at a density of 5 x lo5 cells per ml for 24 h, diluted to a density of 3.5-5.0 X lo5 cells per ml and maintained at this density throughout the expression period (3 days, tk locus; 7 days, hprt locus). Mutation plates for the tk locus were established by seeding AHH-1 cells at a density of 10 000 cells per well in 96-well microtiter plates in 0.2 ml of medium 1640 supplemented with 10% heat-inactivated serum and 4 pg/ml trifluorothymidine (TFI). Plates were incubated for 10 days at 37” C in a 5% CO,, 100% humidified atmosphere, at which time 22 ~1 of complete medium containing 40 pg/ml TFT was added to

S.M. Morris et al. /Mutation Research 329 (1995) 79-96

the individual wells. The plates were evaluated microscopically for the number of positive (normal growth mutants) wells. A well was classified as positive when the clone was a minimum of 1.0 mm in diameter. The clones covered a minimum of 25% of the well bottom and either appeared as a solid mass of cells or had a ‘lacy’ configuration. The plates were returned to the incubator for an additional 10 days and the number of additional positive (slow growth mutants) wells determined microscopically. The wells were classified as positive when the above criteria were met. For the hprt locus, 20000 cells in 0.2 ml of complete medium containing 0.6 pg/ml of 6-TG were seeded into each well of a 96-well plate (Costar, MA). These plates were maintained undisturbed for 13 days and then evaluated microscopically for the presence of positive wells (minimum of 1.0 mm in diameter). Plates were established in triplicate for each concentration of chemical. Plating efficiencies (day 3 and day 7) were determined by seeding 2 cells in 0.2 ml of complete medium without the selective agent into each well of a 96-well plate. Duplicate plates were established for each concentration of mamsa. The plates were evaluated microscopically after 13 days of culture for the number of positive wells. A well was considered positive when the clone was a minimum of 0.25 mm in diameter. Mutation frequencies were determined by the methods of Furth et al. (1981) and plating efficiencies by the methods of Crespi et al. (1985). Cell viability assays In the mutagenesis experiments, exposure to m-amsa resulted in decreased cell numbers at the end of the exposure period; in addition to cell death during the expression period. In order to determine the manner of cell death, AHH-1 cells were exposed with and without serum to increasing concentrations of m-amsa (0.0, 1.0, 10.0, 50.0, 100.0, or 250.0 ng/ml) for 24 h. Aliquots of cells (1 X lo6 cells per concentration) were removed at intervals (-24 h, - 18 h, - 6 h and 0 h) during the exposure period and stained with PI and FDA (see below for protocol). At the end of the exposure period, the m-amsa was removed and the cells seeded at a density of 5 x lo5 cells per

83

ml in complete medium. Aliquots of cells (1 X lo6 cells per concentration) were removed at intervals after exposure (0 h, + 6 h, + 18 h, +24 h, +30 h, +42 h, +48 h, +72 h and -t-168 h) as a means of determining the cell death pathways during a time equivalent to the expression period for the mutagenesis assays. The cells were pelleted by centrifugation, the supernatant removed, and the cells resuspended in 5.0 ml of complete medium with 100 ~1 of 1.0 mg/ml PI staining solution and 200 ~1 of 1 pg/ml FDA staining solution. The cell suspension was incubated at 37” C for 10 min, the cells pelleted by centrifugation, the supernatant removed and the cells resuspended in 1 x PBS, pH 7.4. The cells were maintained under an aluminum foil light shield on ice until FCM analysis. Flow cytometry

Individual samples were analyzed on FACScan (Becton-Dickinson, San Jose, CA) flow cytometer with an argon ion laser tuned to 488 nm. The forward scatter (FSC) signal and the side scatter (SSC) signal were each measured in linear mode with an amplification of 1.5 for FSC and 1.0 for SSC. Both FL1 and FL2 fluorescence were measured in logarithmic amplification, each on a 4decade log scale. 25000 cells were evaluated for each sample utilizing the Consort 30 software (Becton-Dickinson) at a throughput rate of 8001000 cells per second. Electronic compensation (%FLl - %FL2 and %FL2 - %FLl) eliminated cross-signal detection. Individual regions and the percentages of cells within each of the regions were defined in post-acquisition analysis using the Lysis-II software (Becton-Dickinson). Each sample within an individual experiment was analyzed with the same regions to minimize the effect of variation in region size from sample to sample on the percentages of cells within the regions. Statistical methods Induction of mutations at the tk locus and at the hprt locus. The data from the mutation assay

experiments were analyzed using a two-way analysis of variance (ANOVA) with m-amsa concentration and the presence or absence of serum as

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main effects for each locus. The data obtained for each locus (tk-normal growth; &-slow growth; hart) were analyzed separately. A preliminary analysis indicated a significant serum x concentration interaction (p = 0.0043) for the tk-normal growth data. The model for these data included a serum X concentration term. The Dunnett t-test was used to test if any concentration groups were statistically different from the control group. A one-sided test with a significance level of 5% was used for each test. Cell viability assays. The cell viability data were analyzed as previously described (Morris et al., 1994a). Briefly, logistic regression analysis was used to determine the effect of m-amsa concentration and the presence or absence of serum on the proportion of cells within the region of interest. The proportion of cells was determined as the number of cells within one region to the total number of cells analyzed. The test that coefficient p was equal to zero was performed to determine the effects of m-amsa concentration and the presence or absence of serum. Because cells in the same pool of cells tend to have a more similar response than cells from different pools (McCullagh and Nelder, 1989), the quasi-likelihood model was used in fitting the logistic model in order to account for expected similarities within the same pool of cells.

z

Normal-Growth

z

Slow-Growth

___A__

ik f4

k’ti

,’

,I’

Dose @g/ml)

Fig. 1. Induction of mutations in AHH-1 cells exposed to Pn-amsa with serum. Mutations at the hprt locus (A) were determined by resistance to 6-thioguanine. Mutations at the tk locus (0, normal growth; q , slow growth) were determined by resistance to trifluorothymidine. Statistical significance was determined by analysis of variance. The mutant fraction in control cultures was 0.81 x lo@’ at the hprt locus, 10.1 x 10m6 at the tk for the normal growth phenotype and 14.1 x 10m6 at the tk for the slow growth phenotype. Plating efficiencies in control cultures were 47.4% at day 3 and 30.2% at day 7. The results are from duplicate experiments. Materials and methods described in the text.

3. Results Mutation induction The mutant fraction in m-amsa-exposed AHH-1 cells was determined both with and without serum during the exposure period and these results are presented in Figs. 1 and 2, respectively. Exposure to increasing concentrations of m-amsa resulted in a statistically significant (p = 0.0012) increase in the total mutant fraction at the tk locus. The recovery of both normal growth tk mutants and slow growth tk mutants increased significantly (normal growth, p = 0.0003;slow growth, p = 0.0014) with increasing concentration, although the magnitude of the increase in slow growth mutants was greater than that for the normal growth mutants. The concentration-de-

4 20

30

‘lo

50

Dose (ng/ml)

2. Induction of mutations in AHH-1 cells exposed to m-amsa without serum. Mutations at the hprt locus (A) were determined by resistance to 6-thioguanine. Mutations at the tk locus (0, normal growth; n , slow growth) were determined by resistance to trifluorothymidine. Statistical significance was determined by analysis of variance. The mutant fraction in control cultures was 1.12~ lop6 at the hprt locus, 7.39x 10V6 at the tk locus for the normal growth phenotype, and 11.5 x 10m6 at the tk locus for the slow growth phenotype. Plating efficiencies in control cultures were 38.0% at day 3 and 31.2% at day 7. The results are from duplicate experiments. Materials and methods described in the text.

S.M. Morris et al. /Mutation

pendent increase in the mutant fraction at the hprt locus was not statistically significant (p = 0.18801, although a subsequent analysis by linear regression revealed a significant increase (p = 0.0306) in the mutant fraction with increasing concentration. In addition, the recovery of 6-TG’ clones was qualitatively similar to the recovery of normal growth TFT’ clones. The Dunnett t-test demonstrated that the overall increase in the mutant fraction at the tk locus and in the frequency of the normal growth tk clones in cultures exposed to 100 ng/ml was significant (p < 0.05) when compared to the mutant fraction for the DMSO control. Statistically significant increases (p < 0.05) in the mutant fraction for slow growth tk mutants were found for those cultures exposed to 10 ng/ml and 100 ng/ml. No significant differences were found in the induction of 6-TG’ clones. Concentration effects on plating efficiencies were significant for each locus (tk, p = 0.0001; hprt, p = 0.0017) by ANOVA. Plating efficiencies were significantly reduced (p < 0.05) at day 3 (tk locus) in those cultures exposed to the 10,50, and 100 ng/ml concentrations when compared to the

0

100 Dose

200

300

(ng/ml)

Fig. 3. Relative plating efficiency in m-amsa-exposed cells. Plating efficiencies were determined at day 3 (tk locus; circles) and day 7 (hprt locus; triangles) after exposure to increasing concentrations of m-amsa. Exposure to m-amsa was conducted both with serum (open symbols) and without serum (closed symbols). The relative plating efficiency was determined relative to the actual plating efficiency of the appropriate control and assigned a value of 100%. Concentration-response curves fitted by eye. Materials and methods described in the text.

Research 329 (1995) 79-96

85

control and at day 7 (hprt locus) in the cultures exposed to the 50 ng/ml and 100 ng/ml concentrations. No positive wells were observed for either locus when cells were exposed to the 250 ng/ml concentration (Fig. 3). A significant effect (p = 0.0127) of serum deprivation on the frequency of normal growth tk mutants was detected in these experiments. Serum deprivation did not significantly affect the frequency of slow growth tk mutants or the frequency of hprt mutants (tk, slow growth, p = 0.1149; hprt, p = 0.2780). At the lower concentrations of m-amsa (1 and 10 ng/ml), differences in the recovery of slow growth tk mutants were minimal (Fig. 1 vs. Fig. 2). However, at the higher concentrations of m-amsa (50 ng/ml in one experiment and 100 ng/ml in both experiments), no mutants were recovered and the plating efficiencies were essentially zero in those cells exposed in serum-free medium. This is reflected in the fact that plating efficiencies at day 3 (tk locus) were significantly lower (p = 0.0427) in those cultures exposed to m-amsa without serum than in those exposed with serum (Fig. 3). Cell viability Three major populations of AHH-1 cells were revealed by the PI/FDA assay, similar to the results of a previous study (Morris et al., 1994a). In that study, the FDAWSPIneg population was comprised primarily of viable cells, although ‘giant cells’ were present when evaluated microscopically after electronic sorting of the individual populations. Cells undergoing apoptosis were present in the FDAnegPIneg population and the FDAnegPIWS population consisted of cells in the later stages of apoptosis and necrosis. These results have recently been confirmed by transmission electron microscopy of representative cells from the individual sorted populations (Morris, S.M., L.J. McGarrity, O.E. Domon et al., . in preparation). A minor population, FDAPoSPIPoS, was also identified which comprised no more than 5% of the population. Experiments are currently in progress to characterize this population. Representative contour plots are presented for control cells and cells exposed to 250 ng/ml of m-amsa immediately after exposure (Fig. 4A,B),

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S.M. Morris et al. /Mutation Research 329 (1995) 79-96

S.M. Morris et al. /Mutation

Research 329 (1995) 79-96

a7

Dose (nglml) Fig. 5. Effect of exposure to m-amsa on cell viability as determined by the FDA/PI assay. Aliquots of cells (1 x lo61 were removed from the cultures during the exposure period at - 18 h, - 6 h and the 0 h time point. The percentages of viable cells (black bars, FDA~SPI”eg), apoptotic cells (white bars, FDAnesPInesl and late apoptotic and necrotic cells (hatched bars, FDAne8PIPOS)were determined by the FDA/PI assay. Individual regions and the percentages of cells within those regions were determined with the Lysis-II (Becton-Dickinson) software. The mean percentage of cells is presented and represents the data from duplicate experiments. Panel A depicts those experiments in which the exposure to m-amsa was conducted in complete medium. Panel B represent the experiments in which the exposure to m-amsa was conducted in serum-free medium.

at 72 h after exposure (Fig. 4C,D) and at 168 h after exposure (Fig. 4E,F). The effect of exposure to increasing concentrations of m-amsa on the percentage of viable cells, cells undergoing apoptosis, and cells in the later stages of apoptosis and necrosis was determined at multiple time points. In addition, the role of serum deprivation on the recruitment of cells into the death pathways was also examined. During the exposure period, the effect of increasing concentration was to reduce the percentage of viable (FDAPoSPIneg) cells in the population. The decrease in the number of viable cells was statistically significant at the -6 h (p < 0.0001)

and 0 .h (t, = 0.0156) time points. The loss in viability was accounted for by a significant, concentration-dependent increase in the number of cells in the late apoptotic and necrotic cell (FDAnegPIpOS) population (- 18 h, p = 0.0018; -6 h, p < 0.0001; 0 h, p < 0.0001). The effect of increasing concentration of m-amsa on cell viability during the exposure period is shown in Fig. 5A (with serum) and Fig. 5B (without serum). During the expression period, a statistically significant decrease in the proportion of viable cells (FDAPoSPIneg) was found at all time points (0 h, p = 0.0156; 6 h, p = 0.0009; 18 h, p < 0.0001; 24 h, p < 0.0001; 30 h, p < 0.0001; 42 h, p <

Fig. 4. The effect of exposure to m-amsa on the percentage of viable, apoptotic contour plots are presented for cultures exposed to the DMSO control (panels Three time points were selected for display, 0 h (panels A and B), 72 h (panels FL 1 fluorescence (green, FDA) is plotted on the X-axis and FL 2 fluorescence represents 25000 cells. The contour plots were constructed with the Lysis-II algorithm and a 1% threshold.

and necrotic cells in the population. Representative A, C, and El and to 250 ng/ml (panels B, D, and F). C and D) and 168 h (panels E and F) after exposure. (red, PI) is plotted on the Y-axis. Each contour plot software (Becton-Dickinson) with a 1 x smoothing

88

S.M. Morris et al. /Mutation Research 329 (1995) 79-96

1

io

50

Irn

1%

0

1

Dose (nglml) Fig. 6. Effect of exposure to m-amsa in complete medium on the viability of AHH-1 cells. Aliquots of cells (1 X lo? were removed from the cultures at intervals during the post-exposure period and classified as viable, apoptotic or necrotic by the FDA/PI assay. Individual regions and the percentage of cell within each region (black bars, FDAPOSPInes, viable; white bars, FDAnesPIneg, apoptotic; hatched bars, FDA negPIPOS , late apoptotic and necrotic1 were determined with Lysis-II software. The data presented are from two replicate experiments.

0.0001; 48 h, p < 0.0001; 72 h, p < 0.0001; 168 h, p = 0.0007). Although cells were found in the apoptotic population (FDAnegPIneg), the loss in

viability could be accounted for a concentrationdependent increase in the percentage of cells undergoing necrosis or in the later stages of

Fig. 7. Effect of exposure to m-amsa in serum-free medium on the viability of AHH-1 cells. The percentage of viable (black bars, FDA~PIneg), apoptotic (white bars, FDAnesPInea) and late apoptotic and necrotic cells (hatched bars, FDAnegPIWS) was determined by the FDA/PI assay at intervals during the post-exposure periods. Graphics and gating analysis were performed with the Lysis-II (Becton-Dickinson) software. The data presented are from two replicate experiments.

SM. Morris et al. /Mutation Research 329 (199.5) 79-96

apoptosis (FDAnegPIPOS) at the 0 h (p < 0.0001) and 6 h (p < 0.0001) time points. At 18 h, however, the loss in viability related best to the significant, concentration-dependent increase of cells in the FDAnegPIneg (apoptotic) population which was observed at each of the subsequent time points (18 h, p = 0.0008; 24 h, p = 0.0002; 30 h, p = 0.0001; 42 h, p < 0.0001; 48 h, p = 0.0000; 72 h, p < 0.0001; 168 h, p < 0.0001). At 18 h, cells undergoing apoptosis predominated in those cultures exposed to 50 ng/ml, 100 ng/ml and 250 ng/ml of m-amsa and this effect was even more pronounced at the later time points. Cells either in the later stages of apoptosis or undergoing necrosis were present in these populations; however, the number of cells in this population was substantially lower than in the apoptotic population. The percentage of cells in the FDAnegPIPoS (late stage apoptotic and necrotic) population increased with increasing concentration from 18 h until 72 h and then decreased at 168 h. Exposure of AHH-1 cells to m-amsa without serum resulted in an overall decrease in the percentage of viable (FDAPoSPIneg) cells. An increase in the proportion of cells in the FDAnegPIPoS region accounted for the loss of viability due to serum withdrawal during the 24-h exposure to m-amsa. The loss in viability was significant at the - 18 h ( p = 0.0330) and - 6 h (p < 0.0001) time points as was the increase in the FDAnegPIPoS (late stage apoptotic and necrotic) population (-24 h, p < 0.0001; - 18 h, p < 0.0001; - 6 h, p < 0.0001). The loss of viability due to serum deprivation was also evident during the expression period when cells were cultured in complete medium. The decrease in the viable cell population was statistically significant from 6 h (p = 0.0312) to 18 h (p = 0.0381). At 18 h, a significant (p = 0.0052) increase in the proportion of cells in the FDAnegPIneg (apoptotic) population accounted for the loss in viability. An increase in cells undergoing apoptosis (FDAnegPIneg) due to serum deprivation during the exposure period was evident throughout the post-exposure period. The results for selected time points are presented in Fig. 6 for cells exposed with serum (Fig. 6A, 0 h; Fig. 6B, 18 h; Fig. 6C, 24 h; Fig. 6D, 48 h; Fig. 6E, 72 h; Fig. 6F, 168

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h) and in Fig. 7 for cells exposed without serum (Fig. 7A, 0 h; Fig. 7B, 18 h; Fig. 7C, 24 h; Fig. 7D, 48 h; Fig. 7E, 72 h; Fig. 7F, 168 h).

4. Discussion The phenomenon of apoptosis was described originally as a mechanism for the control of cell numbers during differentiation and development (Kerr, 1971; Kerr et al., 1972). Conceptually, the role of apoptosis has been broadened with the recognition that apoptosis is involved in the destruction of cells with DNA damage. Further, cells in which the integrity of the signal transduction pathways leading to apoptosis has been compromised may escape the signal for apoptotic cell death (Barr and Tomei, 1994). Thus, these experiments were conducted to increase our understanding of the role of apoptosis in the recovery of chemically induced mutants in AHH-1 cells. Our first objective was to determine the pathways for cell death in m-amsa-exposed AHH-1 cells. Although cells underwent necrosis, the primary manner of cell death during the expression period was by apoptosis. The appearance of significant numbers of cells undergoing apoptosis was delayed until approximately 18 h in AHH-1 cells, similar to the delayed apoptosis observed in ethylating agent-exposed AHH-1 cells (Morris et al., 1994a) and cells exposed to both monofunctional and cross-linking alkylating agents (Barry et al., 1990). Exposure of proliferating cells to DNA-damaging agents generally results in either a G, arrest which allows excision repair mechanisms to remove DNA adducts before DNA synthesis or a G, arrest where post-replication repair mechanisms attempt to repair DNA damage before cell division (see reviews of Afshari and Barret, 1993; Weinert and Lydall, 1993). The G, arrest is an important cell-cycle checkpoint at which the signal for either mitosis or apoptosis is given, most probably at the level of the ~34~‘~cyclin B complex (Shi et al., 1994). Evidence supporting the importance of this checkpoint for entrance into the apoptotic death pathways was provided by experiments in which cis-platinum-

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exposed proliferating thymocytes underwent apoptosis in G,. Resting thymocytes with the same level of cis-platinum adducts retained cellular viability (Evans and Dive, 1993; Evans et al., 1994). The delayed apoptosis in m-amsa-exposed AHH-1 cells may be at least partially explained by this mechanism since AHH-1 cells respond to m-amsa withdrawal by the completion of S-phase and an arrest in G, (Morris, S.M., L.J. McGarrity, O.E. Domon et al., Cell proliferation and programmed cell death in AHH-1 human lymphoblastoid cells exposed to m-amsa, in preparation), similar to the G, arrest observed in both human and hamster cell lines exposed to topo-II inhibitors (Barry et al., 1990; Zucker et al., 1991; Del Bino and Darzynkiewicz, 1991; Poot et al., 1992). Not all cell lines respond to DNA-damaging agents with a G, arrest and delayed apoptosis. In HL-60 cells which constitutively overexpress c-myc (Collin and Groudine, 1982), death by apoptosis occurs immediately after the withdrawal of mamsa in S-phase cells (Gorczyca et al., 1993b). HL-60 and other cells which overexpress c-myc are sensitive or ‘primed’ to the apoptosis-inducing effects of growth factor withdrawal (Dive and Wyllie, 1993). In these cells, the pathways for apoptotic cell death may be constitutively upregulated, including the synthesis of the apoptosis-associated endonuclease (Wyllie, 1992). The immediate degradation of the DNA in HL-60 cells exposed to topo-I and topo-II inhibitors, as indicated by in situ end-labelling, may reflect the constitutive synthesis of the endonuclease (Gorczyca et al., 1993b). The need for activation of the endonuclease and perhaps, upregulation of other components of the apoptosis pathways may also provide an explanation for the delayed death by apoptosis in our experiments. Alternatively, the inhibition of DNA synthesis due to the persistence of the cleavable complexes (Fox and Smith, 1990) may slow or block the progression of cells towards the G, checkpoint and subsequent signalling for apoptosis. Although the induction of apoptosis was delayed, the signal for cell death most probably occurred during the initial S-phase of exposure. Pretreatment of cells with the DNA replication

inhibitor, aphidicolin, partially protected cells against the cytotoxicity of the topo-I inhibitor, camptothecin, and the topo-II inhibitors, VP-16 and m-amsa. The formation of the cleavable complex was not affected and equal numbers of protein-linked single- and double-strand breaks were detected (Helm et al., 1989; Hsiang et al., 1989). The aphidicolin-induced increase in cell survival, independent of the level of protein-linked break formation, led to the hypothesis that the blockage of replication fork progression may signal cell death (Holm et al., 1989; Hsiang et al., 1989). Support for this model was found in experiments (Charcosset et al., 1985; Chow and Ross, 1987; Schneider et al., 1989) in which cyclohexamide completely inhibited the toxicity of topoisomerases, but not the formation of protein-linked strand breaks. In addition, compounds that do not stabilize the cleavable complex, but apparently distort the superhelicity of the DNA, e.g., fostrecin (Boritzki et al., 1988) and the bis(2,6-dioxopiperazine) derivatives (Onishi et al., 1994), are effective inducers of apoptosis. Experiments were also conducted to determine if cells with DNA damage were able to survive and proliferate under conditions where apoptotic cell death was induced. In these experiments, exposure to m-amsa resulted in a concentration-dependent increase in the mutant fraction at the tk locus, primarily due to the recovery of mutants with the small colony or slow growth phenotype. Molecular and cytological analyses have revealed that the differences in the recovery of mutants between the tk and the hprt locus are due to the presence of tk mutants with the small colony or slow growth phenotype which represent cells with large, intergenic deletions, cells with chromosome rearrangements, or, to a lesser extent, cells that have undergone somatic recombination (Hozier et al., 1982; Yandell et al., 1986; Applegate et al., 1990). The recovery of tk mutants with the small colony phentoype, combined with the studies that indicate that m-amsa is a potent clastogen (Backer et al., 1990; Doerr et al., 19891, is consistent with the presence of chromosomal damage involving the tk locus. Although confirmation of the nature of the DNA damage will require molecular analysis, these results do

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provide evidence that the TFT’ and 6-TG’ clones are derived from cells with DNA damage. These results must be interpreted with respect not only to the current models of mutagenesis, but also to our understanding of the mechanisms of apoptosis. Inherent in the predominant model that addresses the increased recovery of mutants at the tk locus compared to the hprt locus is the concept that mutations that extend beyond the target (tk) gene remain viable due to the presence of linked essential genes on the homologous chromosome (DeMarini et al., 1989). Intergenic mutations at the hemizygous hprt locus may not be recovered due to deletions of genes that confer viability. This model provides an explanation not only for the increased mutation frequency at the tk locus in mouse lymphoma cells heterozygous at the tk locus by clastogenic agents (Moore et al., 1985a,b; DeMarini et al., 1989; Evans et al., 1986), but also offers an explanation for the low recovery of mutations induced at the tk locus in L5178Y cells that are monosomic for chromosome 11 which carries the tk gene (Evans, 1994). Our results, however, suggest that suppression of apoptosis may also be a factor in the viability of cells with multilocus deletions. That is, either pre-existing or concomitant damage to those genes controlling the cell death pathways may block apoptosis and damaged cells are not allowed to die. In the mouse lymphoma cell line, a likely candidate is the murine p53 gene which maps to chromosome 11 (Rotter et al., 1984) as does the tk gene (Hozier et al., 1987). If the ~53 gene is heterozygous and the wild-type (wt) allele is located on the same chromosome as the wt tk allele, then a terminal deletion with breakpoints between the ~53 gene and the centromere would result in a deleted chromosome 11 and an acentric fragment. The acentric fragment, containing both the wt tk and ~53 alleles, would be lost in subsequent cell divisions and result in loss of heterozygosity for both genes. The p53-dependent pathway for apoptosis would be blocked and cells with this mutant phenotype would then survive and proliferate. It is more difficult to extend this hypothesis to human cells in which the ~53 gene (17~13.1, Miller et al., 1986) and the tk gene (17q23-25,

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Human Gene Mapping 7, 1984) map to chromosome 17, but are separated by the centromere. However, several genes that may play a role in apoptosis are located on chromosome 17q. These include c-erbB-2 which maps to 17q21 (Fukushige et al., 1986) and is overexpressed in certain human malignancies (Yamamoto et al., 1986). Interestingly, the mouse homologue to this gene maps to chromosome 11 (Buchberg et al., 19911, the location of ~53 and tk, and may serve a similar function in that species. Also to be considered is the BRCAl gene, important in the etiology of early-onset familial breast cancer, which linkage analyses (Hall et al., 1992; Chamberlin et al., 1993) and molecular analyses (Saito et al., 1993; O’Connell et al., 1994) indicate is located on 17q21. Chromosome transfer experiments (Chen et al., 1994; Theile et al., 1994) also present evidence for additional tumor suppressor genes on both chromosome 17~ and 17q with roles in cellular viability. Alternatively, mutation of the ~53 gene may play a role in the suppression of apoptosis in human cell tk mutants, but it is due to multiple mutagenic events, rather than to a single deletion. This concept, that coincident mutations can affect cell survival and proliferation in tk mutants, is supported by the results of Li et al. (1992) in which tk mutants with the slow growth pheotype from TK6 cells were found to have point mutations, rather than chromosomal rearrangements or intergenic deletions. It was suggested that since a point mutation within the tk gene should not affect the growth characteristics of the mutant clone, the slow growth phenotype was due to additional mutations within the genome. Further, molecular analysis of tk mutants revealed a higher incidence of coincident mutations in minisatellite loci (Li et al., 1992) and microsatellite loci (Li et al., 1994) throughout the genome than was expected based on the mutant fraction at the tk locus. That a pre-existing mutation may affect the viability of damaged cells is suggested by studies in which the overexpression of bcl-2 blocks the bcl-2-dependent apoptosis pathway in maturing T-cells and B-cells (McDonnell et al., 1989, 1990; Tsujimoto, 1989a), or in cells exposed to DNA-

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damaging agents, including inhibitors of topo-I and topo-II (Fanidi et al., 1992; Miyashita and Reed, 1992; Kondo et al., 1994). Gverexpression may be the result of either a chromosomal rearrangement such as the t(14;18) observed in follicular lymphoma, the insertion of the bcl-2 gene and its subsequent upregulation, or, perhaps more relevant to the AHH-1 cell line, transformation by the Epstein-Barr virus (EBV). B-cells, derived from Burkitt lymphoma, which express the full EBV latent genome are resistant to apoptotic cell death under low serum conditions in contrast to those cells which express only the EBNA-1 nuclear antigen and undergo apoptosis with low serum (Gregory et al., 1991). Transfection of the individual EBV latent genes into B-cells indicates that the protective effect is due to the expression of the LMP-1 gene, which, in turn, upregulates bcl-2 expression (Henderson et al., 1991). Further, differences in the cell death rates between cells transfected with the control vector and cells transfected with the bcl-2 vector cannot be accounted for by the level of the initial DNA damage since transfection with the bcl-2 gene did not affect the rates of formation of strand breaks after exposure to inhibitors of thymidylate synthetase (Fisher et al., 1993) or etoposide (Kamesaki et al., 1993). Also of interest were our results in which the exposure of AHH-1 cells to m-amsa without serum resulted in the lowered recovery of mutant cells when compared to cells exposed with serum. The increased rate of apoptosis in the serum-deprived cultures is consistent with a shift in the cellular signalling pathways in favor of apoptosis rather than proliferation. This may be due to the deprivation of growth factors such as the interlet&ins, including IL-3 and IL-6 which modulate the recruitment of cells into the apoptosis pathways (Collins et al., 1992; Yonish-Rouach et al., 1991). In non-transformed cells, serum deprivation results in the entrance of cells into the quiescent or G, state and i’s accompanied by the downregulation of c-myc (Waters et al., 1991). In cells in which the expression of the c-myc protein has been upregulated, cells respond to serum deprivation by undergoing apoptosis (Evan et al., 19921, consistent with the hypothesis of Dive and

Research 329 (1995) 79-96

Wyllie (1993) which suggests that the expression of c-myc ‘primes’ cells for apoptosis. Collins et al. (1992) suggest that IL-3, which protects murine bone marrow cells from the apoptosis-inducing effects of ionizing radiation, etoposide, or cisplatin, may permit cells to progress to a G, checkpoint important in monitoring DNA repair. In other cell lines, the apoptosis-inducing effects of serum withdrawal could be countered with increased expression of the bcl-2 protein (Tsujimoto, 1989a; Milner et al., 1992). In studies in which DNA damage has been measured, modulation of the signalling pathways has not affected the initial level of DNA damage or its repair (Collins et al., 1992). Thus, the death or survival (and subsequent recovery) of mutant cells may depend, at least partially, on factors other than the level of DNA damage. In summary, we have demonstrated that death by apoptosis accounts for the loss of viability in AHH-1 cells exposed to m-amsa. However, the recovery of mutant cells, primarily tk mutants with the slow growth phenotype that is thought to reflect chromosomal mutations, is consistent with the survival and proliferation of damaged cells. This may indicate that the ability to recover mutant cells and the determination of the mutant fraction is influenced, at least partially, by the ability of damaged cells to either suppress the signals for apoptotic cell death or escape from the apoptotic death pathways. The recognition that cell death by apoptosis is an ordered event under genetic control leads to the suggestion that an increased understanding of the mechanisms of apoptosis will also provide insight into the mechanisms of mutagenesis and the relationship between mutagenesis and carcinogenesis.

Acknowledgements The authors would like to acknowledge Dr. David DeMarini (USEPA) for the generous gift of m-amsa, in addition to many helpful comments and suggestions during the course of these experiments. In addition, the authors would like to thank Dr. Bruce Penman (Gentest) for suggesting the methodology for the discrimination of

S. M. Morris et al. /Mutation Research 329 (1995) 79-96

tk mutant phenotypes, Dr. Ben Aidoo (NCTR) for his advice and guidance on the mutagenesis assays, and Dr. M. Manjanatha (NCTR) for his editorial comments on the manuscript.

References Afshari, C.A. and J.C. Barret (1993) Cell cycle controls: Potential targets for chemical carcinogens? Environ. Health Perspect., 101 (~51, 9-14. Applegate, M.L., M.M. Moore, C.B. Broder, A. Burrell, G. Juhn, K.L. Kasweck, P.F. Lin, A. Wadhams and J.C. Hozier (1990) Molecular dissection of mutations at the heterozygous thymidine kinase locus in mouse lymphoma cells, Proc. Natl. Acad. Sci. USA, 87, 51-55. Backer, L.C., J.W. Allen, K. Harrington-Brock, J.A. Campbell, D.M. DeMarini, C.L. Doerr, D.R. Howard, A.D. Kligerman and M.M. Moore (1990) Genotoxicity of inhibitors of DNA topoisomerase I (camptothecin) and II (m-amsa) in vivo and in vitro, Mutagenesis, 5, 541-547. Barr, P.J. and L.D. Tomei (1994) Apoptosis and its role in human disease, Bio/Technology, 12, 487-493. Barry, M.A., CA. Behnke and A. Eastman (1990) Activation of programmed cell death (apoptosis) by cisplatin, other anticancer drugs, toxins and hyperthermia, Biochem. Pharmacol., 40, 2353-2362. Beranek, D.T., CC. Weiss and D.H. Swenson (1980) A comprehensive quantitative analysis of methylated and ethyiated DNA using high-pressure liquid chromatography, Carcinogenesis, 1, 595-606. Boritzki, T.J., T.S. Wolfard, J.A. Besserer, R.C. Jackson and D.W. Fry (1988) Inhibition of type II topoisomerase by fostriecin, Biochem. Pharmacol., 37, 4063-4068. Bronstein, S.M., J.E. Cochrane, T.R. Craft, J.A. Swenberg and T.R. Skopek (1991) Toxicity, mutagenicity, and mutational spectra of N-ethyl-N’-nitrosourea in human cell lines with different DNA repair phenotypes, Cancer Res., 51, 5188-5197. Buchberg, A.M., J.J. Moskow, M.S. Buckwalter and S.A. Camper (1991) Mouse chromosome 11, Mammal. Genome, 1, S158-191. Chamberlin, J.S., M. Boehnke, T.S. Frank, S. Kiousis, J. Xu, S.W. Guo, E.R. Hauser, R.A. Norum, E.A. Helmbold, D.S. Markel, S.M. Keshavarzi, C.E. Jackson, K. Calzone, J. Garber, F.S. Collins and B.L. Weber (1993) BRCAl maps proximal to D17S579 on chromosome 17q21 by genetic analysis, Am. J. Human Genet., 52, 792-798. Charcosset, J.Y., J.P. Bendirdjian, M.F. Lantieri and A. Jacquemin-Sabion (1985) Effects of 9-OH-ellipticine on cell survival, macromolecular syntheses, and cell cycle progression in sensitive and resistant Chinese hamster lung cells, Cancer Res., 45, 4229-4236. Chen, P., N. Ellmore and B. Weissman (1994) Functional evidence for a second tumor suppressor gene on human chromosome 17, Mol. Cell. Biol., 14, 534-542.

93

Chow, KC. and W.E. Ross (1987) Topoisomerase-specific drug sensitivity in relation to cell cycle progression, Mol. Cell Biol., 7, 3119-3123. Clarke, A. R., C.A. Purdle, D.J. Harrison, R.G. Morris, C.C. Bird, M.L. Hooper and A.H. Wyllie (1993) Thymocyte apoptosis induced by p53-dependent and independent pathways, Nature, 362, 849-852. Clive, D. (1987) Historical overview of the mouse lymphoma t/c + / - mutagenicity assay, in: M. Moore, D. DeMarini, F.J. de Serres and K.R. Tindall (Eds.), Banbury Report 28: Mammalian Cell Mutagenesis, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp. 25-36. Collin, S. and M. Groudine (1982) Amplification of endogenous myc-related DNA sequences in a human myeloid leukaemia cell line, Nature, 298, 679-681. Collins, M.K.L., J. Marvel, P. Malde and A. Lopez-Rivas (1992) Interleukin-3 protects murine bone marrow cells from apoptosis induced by DNA damaging agents, J. Exp. Med., 176, 1043-1051. Crespi, C.L., C.G. Ryan, G.M. Seixas, T.R. Turner and B.W. Penman (1985) Tests for mutagenic activity using mutation assays at two loci in the human lymphoblast cell lines, TK6 and AHH-1, Prog. Mutation Res., 5, 497-516. Del Bino, G. and Z. Darzynkiewicz (1991) Camptothecin, teniposide, or 4’-(9-acridinylamino)-3-methanesulfon-maniside, but not mitoxantrone or doxorubicin, induces degradation of nuclear DNA in the S phase of HL-60 cells, Cancer Res., 51, 1165-1169. Del Bino, G., J.S. Skierski and Z. Darzynkiewicz (1991) The concentration-dependent diversity of effects of DNA topoisomerase I and II inhibitors on the cell cycle of HL-60 cells, Exp. Cell Res., 19.5,485-491. DeMarini, D., CL. Doerr, M.K. Meyer, K.H. Brock, J. Hozier and M.M. Moore (1987) Mutagenicity of m-amsa and o-amsa in mammalian cells due to clastogenic mechanism: possible role of topoisomerase, Mutagenesis, 2, 349-355. DeMarini, D., H.E. Brockman, F.J. de Serres, H.H. Evans, L.F. Stankowski and A.W. Hsie (1989) Specific-locus mutations in eucaryotes (especially mammalian cells) by radiation and chemicals: a perspective, Mutation Res., 220, 11-29. Dive, C. and A.H. Wyllie (1993) Apoptosis and cancer chemotherapy, in: J.A. Hickman and T.T. Tritton (Eds.), Frontiers in Pharmacology: Cancer Chemotherapy, Blackwell Scientific, Oxford, pp. 21-56. Doerr, C.L., K. Harrington-Brock and M.M. Moore (1989) Micronucleus, chromosome aberration, and small-colony TK mutant analysis to quantitate chromosomal damage in L5178Y mouse lymphoma cells, Mutation Res., 222, 191203. Donehower, L.A. and A. Bradley (1993) The tumor suppressor ~53, Biochim. Biophys. Acta, 1155, 181-205. Evan, G., A.H. Wyllie, C.S. Gilbert, T.D. Littlewood, H. Land, M. Brooks, CM. Waters, L.Z. Penn and D.C. Hancock (1992) Induction of apoptosis in fibroblasts by c-myc protein, Cell, 69, 119-128. Evans, CA., P.J. Gwen-Lynch, A.D. Whetton and C. Dive

94

S.M. Morris et al. /Mutation Research 329 (1995) 79-96 (1993) Activation of the Abelson tyrosine kinase activity is

associated with suppression of apoptosis in hemopoietic cells, Cancer Res., 53, 1735-1738. Evans, D.L. and C. Dive (1993) Effects of cisplatin on the induction of apoptosis in proliferating hepatoma cells and nonproliferating immature thymocytes, Cancer Res., 53, 2133-2139. Evans, D.L., M. Tilby and C. Dive (1994) Differential sensitivity to the induction of apoptosis by cisplatin in proliferating and quiescent immature rat thymocytes is independent of the levels of drug accumulation and DNA adduct formation, Cancer Res., 54, 1.596-1603. Evans, H.H. (1994) The prevalence of multilocus lesions in radiation-induced mutants, Radiat. Res., 137, 131-144. Evans, H.H., J. Mencl, M.F. Horng, M. Ricanti, C. Sanchez and J. Hazier (1986) Locus specificity in the mutability of L5178Y mouse lymphoma cells: The role of multilocus lesions, Proc. Natl. Acad. Sci. USA, 83, 4379-4383. Fanidi, A., E.A. Harrington and G.I. Evan (1992) Cooperative interaction between c-myc and bd-2 protooncogenes, Nature, 359, 554-556. Fisher, T.C., A.E. Milner, CD. Gregory, A.L. Jackman, G.W. Aherne, J. Hartley, C. Dive and J.A. Hickman (1993) bcl-2 modulation of apoptosis induced by anticancer drugs: resistance to thymdiylate stress is independent of classical resistance pathways, Cancer Res., 53, 3321-3326. Fox, M.E. and P.J. Smith (1990) Long-term inhibition of DNA synthesis and the persistence of trapped topoisomerase-II complexes in determining the toxicity of the antitumor intercalators mAMSA and mitoxantrone, Cancer Res., 50, 5813-5818. Fukushige, S.I., K.I. Matsubara, M. Yoshida, M. Sasaki, T. Suzuki, K. Semba, K. Toyoshima and T. Yamamoto (1986) Localization of a novel u-erbB-related gene, c-erbB-2, on human chromosome 17 and its amplification in a gastric cancer cell line, Mol. Cell. Biol., 6, 955-958. Furth, E., W.G. Thilly, B.W. Penman, H.L. Liber and W.M. Rand (1981) Quantitative assay for mutation in diploid human lymphoblasts using microtiter plates, Anal. Biochem., 110, l-8. Gorczyca, W., J. Gong and Z. Darzynkiewicz (1993a) Detection of DNA strand breaks in individual apoptotic cells by the in situ terminal deoxynucleotidyl transferase and nick translation assays, Cancer Res., 53, 1945-1951. Gorczyca, W., J. Gong, B. Ardelt, F. Traganos and Z. Darzynkiewicz (1993b) The cell cycle related differences in susceptibility of HL-60 cells to apoptosis induced by various antitumor agents, Cancer Res., 53, 3186-3192. Gorczyca, W., M.R. Melamed and 2. Darzynkiewicz (1993~) Apoptosis of S-phase HL-60 cells induced by topoisomerase inhibitors: Detection of DNA strand breaks by flow cytometry using the in situ nick translation assay, Toxicol. Lett., 67, 249-258. Gregory, C.D., C. Dive, S. Henderson, C.A. Smith, G.T. Williams, J. Gordon and A.B. Rickinson (1991) Activation of Epstein-Barr virus latent genes protects human B cells from death by apoptosis, Nature, 349, 612-614. Hall, J.M., L. Friedman, C. Guenther, M.K. Lee, J.L. Weber,

D.M. Black and M.C. King (1992) Closing in on a breast cancer gene on chromosome 17q, Am. J. Hum. Genet., 50, 1235-1242. Heflich, R.H., D.T. Beranek, R.L. Kodell and S.M. Morris (1982) Induction of mutations and sister-chromatid exchanges in Chinese hamster ovary cells. Relationship to specific DNA adducts, Mutation Res., 106, 147-161. Henderson, S., M. Rowe, C. Gregory, D. Croom-Carter, F.Wang, R. Longnecker, E. Kieff and A. Rickinson (1991) Induction of bd-2 expression by Epstein Barr virus latent membrane protein 1 protects infected B cells from programmed cell death, Cell, 65, 1107-1115. Hickman, J.A. (1992) Apoptosis induced by anticancer drugs, Cancer Metast. Rev., 11, 121-139. Hockenberxy, D., G. Nunez, C. Millman, R.D. Schreiber and S.J. Korsemeyer (1990) bcl-2 is an inner mitochondrial membrane protein that blocks programmed cell death, Nature, 348, 334-336. Holm, C., J.M. Covey, D. Kerrigan and Y. Pommier (1989) Differential requirement of DNA replication for the cytotoxicity of DNA topoisomerase I and II inhibitors in Chinese hamster DC3F cells, Cancer Res., 49, 6365-6368. Hozier, J., J. Sawyer, M. Moore, B. Howard and D. Clive (1981) Cytogenetic analysis of the L5178/TK+ /--* TK-/mouse lymphoma mutagenesis assay system, Mutation Res., 84, 169-181. Hozier, J., J. Sawyer, D. Clive and M. Moore (1982) Cytogenetic distinction between the TK+ and TK- chromosome in the L5178 TK+ \ - mouse lymphoma mutagen assay system, Mutation Res., 105, 451-456. Hozier, J., M. Applegate, N. deVassa1 and J. Sawyer (1987) Localization of the cytoplasmic thymidine kinase (X-1) on mouse chromosome 11 by in situ molecular hybridization and comparison of human and mouse mapping sites at high cytogenetic resolution, Cytogenet. Cell Genet., 46, 631-640. Hsiang, Y.H., M.G. Lihou and L.F. Liu (1989) Arrest of replication forks by drug-stabilized topoisomerase I-DNA cleavable complexes as a mechansim of cell killing by camptothecin, Cancer Res., 49, 5077-5082. Human Gene Mapping 7 (1984) Cytogenet. Cell Genet., 37, 3-393. Jaxel, C., G. Tadou, C. Portemer, G. Mirambeau, J. Panijel and M. Duget (1988) Topoisomerase inhibitors induce irreversible fragmentation of replicated DNA in concanavalin A stimulated splenocytes, Biochemistry, 27, 9599. Kamesaki, S., H. Kamesaki, T.J. Jorgenson, A. Tanizawa, Y. Pommier and J. Cossman (1993) bcl-2 protein inhibits etoposide-induced apoptosis through its effects on events subsequent to topoisomerase II-induced DNA strand breaks and their repair, Cancer Res., 53, 4251-4256. Kerr, J.F.R. (1971) Shrinkage necrosis: A distinct mode of cellular death, J. Pathol., 105, 13-20. Kerr, J.F.R., A.H. Wyllie and A.R. Currie (1972) Apoptosis: A basic biological phenomenon with wide ranging implications in tissue kinetics, Br. J. Cancer, 26, 239-257. Kondo, S., D. Yin, T. Morimura and J. Takeuchi (1994) bcl-2

S.M. Morris et al. /Mutation gene prevents induction of apoptosis in L1210 murine leukemia cells by SN-38, a metabolite of the camptothecin derivative CPT-11, Int. J. Oncol., 4, 649-654. Korsemeyer, S.J. (1992a) bcl-2 inititates a new category of oncogenes: Regulators of cell death, Blood, 80, 879-886. Korsemeyer, S.J. (1992b) bcl-2: a repressor of lymphocyte death, Immunol. Today, 13, 285-288. Lane, D.P. (1993) A death in the life of ~53, Nature, 362, 786-787. Li, C.Y., D.W. Yandell and J.B. Little (1992) Molecular mechanisms of spontaneous and induced loss of heterozygosity in human cells in vitro, Somat. Cell Mol. Genet., 18, 77-87. Li., C.Y., D.W. Yandell and J.B. Little (1994) Elevated frequency of microsatellite mutations in TK6 human lymphoblast clones selected for mutations at the thymidine kinase locus, Mol. Cell. Biol., 14, 4373-4379. Liber, H.L., D.W. Yandell and J.B. Little (1989) A comparison of mutation induction at the tk and hprt loci in human lymphoblastoid cells: quantitative differences are due to an additional class of mutations at the autosomal tk locus, Mutation Res., 216, 9-17. Liu, L.F. (19891 DNA topoisomerase poisons as antitumor drugs, Annu. Rev. B&hem., 58, 351-375. Lowe, S.W.. E.M. Schmitt, A.W. Smith, B.A. Osborne and T. Jacks (1993) p53 is required for radiation-induced apoptosis in mouse thymocytes, Nature, 362, 847-848. Marshall, B. and R.K. Ralph (1985) The mechanism of action of mAMSA, Adv. Cancer Res., 44, 267-293. McCullagh, J. and J.A. Nelder (1989) Generalized Linear Model, 2nd edn., Chapman and Hall, London. McDonnell, T.J. (1993) Cell division versus cell death: A functional model of multistep neoplasia, Mol. Carcinogen., 8, 209-213. McDonnell, T.J., N. Deane, F.M. Platt, G. Nunez, U. Jaeger, J.P. McKearn and S.J. Korsemeyer (1989) bcl-2-immunoglobulin transgenic mice demonstrate extended B cell survival and follicular lymphoproliferation, Cell, 57, 79-88. McDonnell, T.J., G. Nunez, F.M. Platt, D. Hockenberry, L. London, J.P. McKearn and S.J. Korsemeyer (1990) Deregulated bcl-2-immunoglobulin transgene expand a resting but responsive immunoglobulin M and D-expressing B-cell population, Mol. Cell. Biol., 10, 1901-1907. Milner, A.E., G.D. Johnson and C.D. Gregory (1992) Prevention of programmed cell death in Burkitt lymphoma cell lines by bcl-2-dependent and -independent mechanisms, Int. J. Cancer, 52, 636-644. Miller, C., T. Mohandas, D. Wolf, M. Prokocimer, V. Rotter and H.P. Koeffler (1986) Human ~53 gene localized to short arm of chromosome 17, Nature, 319, 783-784. Miyashita, T. and J.C. Reed (1992) bcl-2 gene transfer increases relative resistance of S49.1 and WEH17.2 lymphoid cells to cell death and DNA fragmentation induced by glucocorticoids and mutiple chemotherapeutic drugs, Cancer Res., 52, 5407-5411. Moore, M. M., D. Clive, B.E. Howard, A.G. Batson and N.T. Turner (1985a) In situ analysis of trifluorothymidine

Research 329 (1995) 79-96

95

(TIT’) mutants of L5178Y/TK+ / - mouse lymphoma cells, Mutation Res., 151, 147-159. Moore, M.M., D. Clive, J.C. Hozier, B.E. Howard, A.G. Batson, N.T. Turner and J. Sawyer (1985b) Analysis of trifluorothymidine-resistant (TFT’) mutants of L5178Y/TK+/mouse lymphoma cells, Mutation Res., 151, 161-174. Morris, S.M., L.J. McGarrity, O.E. Domon, J.J. Chen, W.G. Hinson, T.J. Bucci, A.R. Warbritton and D.A. Casciano (1994a) The role of programmed cell death in the toxicity of the mutagens, ethyl methanesulfonate and N-ethyl-N’nitrosourea, in AHH-1 human lymphoblastoid cells, Mutation Res., 306, 19-34. Morris, S.M., O.E. Domon, K.B. Delclos, J.J. Chen and D.A. Casciano (1994b) Induction of mutations at the hypoxanthine phosphoribosyl transferase (HPRT) locus in AHH-1 human lymphoblastoid cells, Mutation Res., 310, 45-54. Oberhammer, F., J.W. Wilson, C. Dive, I.D. Morris, J.A. Hickman, A.E. Wakeling, P.R. Walker and M. Sikorska (1993) Apoptotic death in epithelial cells: cleavage of DNA to 300 and/or 50 kb fragments prior to or in the absence of internucleosomal fragmentation, EMBO J., 12. 3679-3684. O’Connell, P., H. Albertson, N. Matsunami, T. Taylor, J.E. Hundley, T.L. Johnson-Pais, B. Reus, E. Lawrence, L. Ballard. R. White and R.J. Leach (1994) A radiation hybrid map of the BRCAl region, Am J. Hum. Genet., 54, 526-534. Onishi, Y., Y. Azuma and H. Kizaki (1994) Bis(2,6-dioxopiperazine) derivatives, topoisomerase inhibitors which do not form a DNA cleavable complex, induce thymocyte apoptosis, Biochem. Mol. Biol. Int., 32, 115-122. Oren, M. (19921 The involvement of oncogenes and tumor suppressor genes in the control of apoptosis, Cancer Metast. Rev., 11, 141-148. Pavlik, E.J., R.C. Flanigan, J.R. van Nagell, M.B. Hanson, E.S. Donaldson, K. Keaton, B. Doss, J. Bartman and D.E. Kenady (1984) Esterase activity, exclusion of propidium iodide, and proliferation in tumor cells exposed to anticancer agents: Phenomenon relevant to chemosensitivity determinations, Cancer Invest., 3, 413-426. Penman, B.W. and C.L. Crespi (1987) Analysis of human lymphoblast mutation assays by using historical negative control data bases, Environ. Mol. Mutagen., 10, 35-60. Poot, M., B. Epe and H. Hoehn (1992) Cell cycle effects of the DNA topoisomerase inhibitors camptothecin and MAMSA in lymphoblastoid cell lines from patients with Fanconi anemia, Mutation Res., 270. 185-189. ROSS, D.D., C.C. Joneckis, J.V. Ordonez, A.M. Sisk, R.K. Wu, A.W. Hamburger and R.E. Nora (1989) Estimation of cell survival by flow cytometric quantification of fluorescein diacetate/propidium iodide viable cell number, Cancer Res., 49. 3776-3782. Rotter, V., D. Wold, D. Pravtcheva and F.H. Ruddle (1984) Chromosomal assignment of the murine gene encoding the transformation-related protein ~53, Mol. Cell Biol., 4, 382-385.

96

S.M. Morris et al. /Mutation Research 329 (1995) 79-96

Saito, H., J. Inazawa, S. Saito, F. Kasumi, S. Koi, S. Sagae, R. Kudo, J. Saito, K. Noda and Y. Nakamura (1993) Detailed deletion mapping of chromosome 17q in ovarian and breast cancers: ~-CM region on 17q21.3 often and commonly deleted in tumors, Cancer Res., 53, 3382-3385. Schneider, E., P.A. Lawson and R.R. Ralph (1989) Inhibition of protein synthesis reduced the cytotoxicity of 4’-(9acridinylamino)methanesulfon-m-anisidide without affecting DNA breakage and DNA topoisomerase II in a murine mastocytoma cell line, Biochem. Pharmacol., 38, 263-269. Shi, L., W.K. Nishioka, J. Th’ng, E.M. Bradbury, D.W. Litchfield and A.H. Greenberg (1994) Premature ~34~~~’ activation required for apoptosis, Science, 263, 1143-1145. Shibuya, M.L., A.M. Ueno, D.B. Vannais, P.A. Craven and CA. Waldren (1994) Megabase pair deletions in mutant mammalian cells following exposure to amsacrine, an inhibitor of DNA topoisomerase-II, Cancer Res., 54, 10921097. Theile, M., S. Hartmann, H. Naundorf, D. Ruess, B. Elbe, H. Krause, W. Deppert, J.C. Barrett and S. Scherneck (1994) Wild-type p53 is not involved in reversion of tumorigenic phenotype of breast cancer cells after transfer of normal chromosome 17, Int. J. Oncol., 4, 1067-1075. Tsujimoto, Y. (1989al Stress-resistance conferred by high level of bcl-2 protein in human B lymphoblastoid cell, Oncogene, 4, 1331-1336. Tsujimoto, Y. (1989b) Gverexpression of the human bcl-2 gene product results in growth enhancement of EpsteinBarr virus immortalized B cells, Proc. Natl. Acad. Sci. USA, 86, 1958-1962. Tsujimoto, Y., J. Cossman, E. Jaffe and CM. Croce (1985) Involvement of the bcl-2 gene in human follicular lymphoma, Science, 228, 1440-1443. Vaux, D.L., S. Cory and J.M. Adams (19881 bcl-2 gene promotes haemopoietic cell survival and cooperates with c-myc to immortalize pre-B cells, Nature, 335, 440-442.

Walker, P.R., C. Smith, T. Youdale, J. Leblanc, J.F. Whitfield and M. Sikorska (1991) Topoisomerase-II-reactive chemotherapeutic drugs induce apoptosis in thymocytes, Cancer Res., 51, 1078-1085. Walker, P.R., L. Kokileve, J. LeBlanc and M. Sikorska (1993) Detection of the initial stages of DNA fragmentation in apoptosis, BioTechniques, 15, 1032-1040. Waters, C., T. Littlewood, D. Hancock, J. Moore and G. Evan (19911 c-myc protein expression in untransformed fibroblasts, Oncogene, 6, 101-109. Weinert, T. and D. Lydall (1993) Cell cycle checkpoints, genetic instability and cancer, Semin. Cancer Biol., 4, 129-140. Wyllie, A.H. (1992) Apoptosis and the regulation of cell numbers in normal and neoplastic tissues: an overview, Cancer Metast. Rev., 11, 95-103. Yamamoto, T., S. Ikawa, T. Akiyama, K. Semba, N. Nomura, N. Miyajima, T. Saito and K. Toyoshima (1986) Similarity of protein encoded by the human c-erbB-2 gene to epiderma1 growth factor receptor, Nature, 319, 230-234. Yandell, D.W., T.P. Dryja and J.B. Little (1986) Somatic mutations at a heterozygous autosomal locus in humans cells occur more frequently by allele loss than by intragenie structural alterations, Somat. Cell Mol. Genet., 12, 255-263. Yonish-Rouach, E., D. Resnitzky, J. Lotem, L. Sachs, A. Kimchi and M. Oren (1991) Wild-type p53 induces apoptosis of myeloid leukaemic cells that is inhibited by interleukin-6, Naiure, 352, 345-347. Zucker, Z.M., D.J. Adams, K.W. Bair and K.H. Elstein (1991) Polyploidy induction as a consequence of topoisomerase inhibition. A flow cytometric assessment, Biochem. Pharmacol., 42, 2199-2208.