* Faculty of Integrated Arts and Sciences, Hiroshima University, Higashi-Hiroshima, 724, Japan; and Gene Expression
Laboratory, The Salk Institute for Biological Studies, La Jolla, California 92037
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Abstract |
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Acentric, autonomously replicating extrachromosomal structures called double-minute chromosomes (DMs) frequently mediate oncogene amplification in human tumors. We show that DMs can be removed from the nucleus by a novel micronucleation mechanism that is initiated by budding of the nuclear membrane during S phase. DMs containing c-myc oncogenes in a colon cancer cell line localized to and replicated at the nuclear periphery. Replication inhibitors increased micronucleation; cell synchronization and bromodeoxyuridine-pulse labeling demonstrated de novo formation of buds and micronuclei during S phase. The frequencies of S-phase nuclear budding and micronucleation were increased dramatically in normal human cells by inactivating p53, suggesting that an S-phase function of p53 minimizes the probability of producing the broken chromosome fragments that induce budding and micronucleation. These data have implications for understanding the behavior of acentric DNA in interphase nuclei and for developing chemotherapeutic strategies based on this new mechanism for DM elimination.
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Introduction |
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THE accumulation of structural and numerical chromosome abnormalities in eukaryotic cells is limited
by coordinating biosynthetic and repair processes
with cell cycle checkpoints (Hartwell and Kastan, 1994).
Mutations in genes involved in these transactions occur
commonly during cancer progression and can greatly elevate the frequencies of base alterations or large-scale
chromosome rearrangements. For example, defects in cell
cycle-control pathways involving the p53 tumor suppressor gene create a permissive environment in which cells
with aneuploidy, chromosome translocations, and gene
amplification arise at high frequency in response to
stresses created by antimetabolites or oncogene overexpression (Livingstone et al., 1992
; Yin et al., 1992
; Denko
et al., 1994
).
The types of aberrant chromosomal structures generated in cells with defective repair and cell cycle control
functions are likely to be constrained by nuclear structure.
For example, chromosomes with very long arms tend to
generate nuclear projections variously referred to as
"blebs" or "buds" (Ruddle, 1962; Lo and Fraccaro, 1974
;
Toledo et al., 1992
; Pedeutour et al., 1994
). A recent study in peas demonstrated that excessive DNA within a single
chromosome arm generated a nuclear projection that was
cut when the cell division plate formed after telophase
(Schubert and Oud, 1997
). Sequences enclosed in such
projections are often detected in micronuclei, suggesting
that projections can be precursors of micronuclei (Toledo et al., 1992
; Pedeutour et al., 1994
), and that the chromosomal sequences they contain can be lost from the nucleus.
These data indicate that a maximum allowable size exists
for each chromosome arm within the nuclei of specific cell
types.
Circular, autonomously replicating DNA fragments
such as double-minute chromosomes (DMs)1 are also frequently generated in cancer cells (Barker, 1982; Cowell,
1982
; Benner et al., 1991
). These structures encode proteins that provide survival advantages in vivo, or resistance
to a variety of chemotherapeutic agents in vitro (Alitalo
and Shwab, 1986
; Wahl, 1989
; Von Hoff et al., 1992
; Brison, 1993
; Shimizu et al., 1994
; Eckhardt et al., 1994
). DMs
replicate using cellular replication origins (Carroll et al.,
1993
), but lacking centromeres, they do not segregate by
the same mechanisms used by chromosomes. Consequently, DMs are lost spontaneously in the absence of selection. Drugs such as hydroxyurea (HU) significantly increase the loss rate of DMs in human and rodent cell lines
(Snapka and Varshavsky, 1983
; Von Hoff et al., 1991
; Von
Hoff et al., 1992
; Eckhardt et al., 1994
; Canute et al., 1996
).
DM elimination results in increased drug sensitivity, reduced tumorigenicity, or differentiation, depending on the
proteins expressed by DM-encoded genes (Snapka and
Varshavsky, 1983
; Snapka, 1992
; Von Hoff et al., 1992
;
Eckhardt et al., 1994
; Shimizu et al., 1994
). Identifying the
mechanisms by which DMs are eliminated could enable
the development of new and more selective chemotherapeutic strategies, since DMs are uniquely found in cancer
cells, and chromosome loss should not be induced by such
treatments.
Like abnormally long chromosome arms, DMs have also
been reported to be preferentially incorporated within micronuclei that are removed from the cell (Von Hoff et al.,
1992; Shimizu et al., 1996
). It is clear that small size alone
does not guarantee selective enclosure of DNA fragments
within micronuclei because a centric minichromosome the
size of a typical DM is effectively excluded from micronuclei (Shimizu et al., 1996
). This observation is consistent with the classical mechanism of micronucleus formation
that involves the enclosure of lagging acentric chromosome fragments as nuclear membranes reform at the end
of mitosis (Heddle and Carrano, 1977
; Heddle et al., 1983
).
Thus, one would expect postmitotic enclosure of DMs
within micronuclei since they typically lack functional centromeres (Levan et al., 1976
). However, DMs appear to
associate with chromosomes or nucleoli, which may enable
most of them to evade such a postmitotic mechanism. The
ability of DMs to "hitchhike" by association with mitotic
chromosomes or nucleoli provides one explanation of why
few micronuclei were detected at the midbody in a cell line
containing numerous DMs (Levan and Levan, 1978
), and
their surprisingly efficient partitioning to daughter cells in
some cell lines (Levan and Levan, 1978
; Hamkalo et al.,
1985
). However, the interphase behavior of normal chromosomes and DMs may differ because DMs lack the centromeres and/or telomeres that position chromosomes in
restricted territories and produce a choreographed set of
chromosome movements during S phase (DeBoni and
Mintz, 1986
; Cremer et al., 1993
). It has neither been determined whether acentric DM-DNA occupies positions
different from chromosomes in interphase, nor whether
this could enable their removal from the nucleus by a budding process like that observed for abnormally long chromosomes (Ruddle, 1962
; Jackson and Clement, 1974
; Lo and Fraccaro, 1974
; Miele et al., 1989
; Toledo et al., 1992
).
DMs provide an excellent model for analyzing the nuclear behavior of multimegabase replicons lacking a centromere and telomeres. We show that DMs can preferentially localize the nuclear periphery, whereas chromosomally amplified sequences occupy a more central position. Although the micronuclei that entrap acentric chromosome fragments have typically been viewed to be generated postmitotically, we provide evidence for a novel micronucleation mechanism that involves the formation of nuclear projections which we refer to as buds. Buds form during S phase and appear to selectively associate with DMs replicating near or at the nuclear periphery. Since micronuclei are indicators of DNA damage and are produced at much higher frequencies in tumor cells than in normal cells, we investigated whether micronucleation frequency is increased in cells with defects in cellular responses to DNA damage. We show that loss of p53 function increases the frequency of micronucleation and enables buds and micronuclei to be produced under conditions expected to lead to chromosome breakage.
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Materials and Methods |
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Cell Culture
Human COLO 320DM (CCL 220) and COLO 320HSR (CCL 220.1) neuroendocrine tumor cells were obtained from American Type Culture Collection (Rockville, MD) and then single cell subclones were obtained by
limiting dilution (Von Hoff et al., 1988). The locations of amplified c-myc
genes to DMs or HSRs were confirmed by fluorescence in situ hybridization (FISH) using c-myc cosmid DNA. The cells were grown in RPMI
1640 medium supplemented with 10% FBS. The WS1 human embryonic
skin fibroblast, obtained from American Type Culture Collection (CRL
1502), was cultured in DME supplemented with 10% heat-inactivated, dialyzed FBS, and 1× MEM nonessential amino acids. WS1-neo and WS1-E6
were kind gifts of S. Linke (National Institutes of Health, Bethesda, MD),
and were generated by infecting WS1 with retroviral vectors expressing
genes encoding either neomycin resistance or both neomycin resistance
and the E6 protein from human papilloma virus 16, respectively (Linke et
al., 1996
). RPE-h (normal human retinal pigmented epithelial cells) and
its neo and E6 derivatives were also kindly provided by S. Linke and the parental cells were obtained from Cell Genesys, Inc. (Foster City, CA). Epithelial cells were cultured in the same way as WS1.
Chemicals
Aphidicolin, 5-bromo-2'-deoxyuridine (BrdU), coumarin (1, 2-benzopyrone), deferoxamine mesylate (desferrioxamine mesylate), DMSO, hydroxyurea, nicotinamide, thymidine, and nocodazole (methyl-[5-{2-thienylcarbonyl}-1H-benzimidazol-2-YL]carbamate) were obtained from Sigma Chemical Co. (St. Louis, MO). Guanazole (3,5-diamino-1,2,4-triazole) was from Aldrich Chemical Co. (Milwaukee, WI). PALA (N-phosphonacetyl-L-aspartate) was provided by the Drug Biosynthesis and Chemistry Branch, Developmental Therapeutics Program, Division of Cancer Treatment, National Cancer Institute (Bethesda, MD).
Cell Cycle Analysis
Cell cycle distribution was analyzed using flow cytometry as described
previously (Yin et al., 1992; Di Leonardo et al., 1994
). Cells treated with
the indicated concentrations of drugs for the indicated times were labeled
with 10 µM BrdU for 30 min. The cells were collected, fixed with 70% ethanol, treated with 0.1 N HCl containing 0.5% Triton X-100 (Mallinckrodt,
Paris, KY), and then followed by boiling for 10 min and rapid cooling to
denature the DNA. The nuclei were then incubated with FITC-conjugated anti-BrdU antibodies (Boehringer Mannheim Biochemicals, Indianapolis, IN) and counterstained with 2 µg/ml of propidium iodide (PI)
containing RNase (200 µg/ml). Samples were analyzed using a Becton
Dickinson FACScanTM (Sparks, MD). 10,000 events were collected for
each sample. Data were analyzed using Sun Display as described previously (Yin et al., 1992
; Di Leonardo et al., 1994
).
Quantification of Micronuclei
Micronuclei containing DM sequences in COLO 320DM cells (see Figs. 2
and 3) were detected by preparing chromosome spreads using standard
hypotonic swelling conditions (Lawce and Brown, 1991), followed by
hybridization with a biotinylated c-myc cosmid probe as described previously (Shimizu et al., 1996
). Total micronuclei (see Fig. 3) were determined by staining chromosome spreads with 4',6-diamidino-2-phenylindole
(DAPI) (Sigma Chemical Co.; 1 µg/ml in VectaShield, Vector Labs, Inc.,
Burlingame, CA). The adherent cells (WS1, RPE-h, and their derivatives)
were grown on coverslips, fixed with cold acetone (
20°C for 5 min) followed by cold methanol (
20°C for 5 min), rehydrated with PBS, and
then stained with DAPI (1 µg/ml in VectaShield). The numbers of total or
DM-enriched micronuclei were scored using 60 or 100× objectives and a
fluorescence microscope equipped with appropriate epifluorescence filters (model Zeiss WL; Carl Zeiss, Inc., Thornwood, NY). The results are
expressed as Frequency of Micronuclei (%) relative to the number of interphase nuclei scored (
1,000 for each point).
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Cell Synchronization
Synchronization was performed as described previously (Stein et al.,
1994). Rapidly growing COLO 320DM cells were first arrested in early S
phase using excess thymidine (2 mM) for 17 h. The cells were then washed
with growth medium, released into growth medium containing 25 µM 2'-deoxycytidine for 12 h (to reverse thymidine toxicity), and then incubated
in 2.5 µg/ml aphidicolin for 17 h to arrest cells as they entered S phase.
The arrested cells were washed with growth medium and then either released into medium lacking drug, or into medium containing nocodazole
(0.4 µg/ml). Cell cycle progression was monitored using incorporation of
[3H]thymidine (Stein et al., 1994
). To monitor the progression through mitosis, a portion (1 ml) of culture was fixed by paraformaldehyde (PFA;
2%) and then stained with DAPI. The frequency of cells in mitosis was
scored using fluorescence microscopy. WS1-E6 cells were synchronized by
seeding them at low density in 15-cm dishes with or without coverslips
(18 × 18 mm). The day after subculture, the medium was removed and
then replaced with medium containing 0.1% FCS, and then followed by
the culture for an additional 48 h. Cells arrested at G0 by serum deprivation were released into growth medium containing 5 µg/ml aphidicolin for
15 h to arrest them at the beginning of S phase. Cells were released into S
phase by replacing the medium with fresh growth medium lacking drug. Progression into S phase was monitored by the incorporation of [3H]thymidine (Stein et al., 1994
). The number of cells that were in S phase just
before release from synchrony was monitored by BrdU-pulse labeling (30 min) followed by confocal examination of the labeling pattern as described in the following section (see Simultaneous Determination of DM
Location and DNA Replication Using FISH and BrdU Incorporation).
Concurrently, coverslips were removed, fixed with acetone and methanol,
stained by DAPI, and then the frequency of micronuclei, nuclear budding,
and mitotic cells were scored as described above.
Terminal Deoxynucleotidyl Transferase-mediated dUTP-Biotin Nick-end Labeling Assay
Terminal deoxynucleotidyl transferase-mediated dUTP-biotin nick-end
labeling (TUNEL) assay was done according to a previously published
procedure (Gavrieli et al., 1992) modified as described below. In brief,
COLO 320DM cells were fixed in 2% PFA (10 min at room temperature)
and then centrifuged onto a glass slide using a cytospin (Salk Institute
Shop) apparatus. The cells were further fixed in cold methanol (
20°C, 5 min) followed by cold acetone (
20°C, 5 min). The slides were rehydrated
in PBS and then equilibrated in reaction buffer (200 mM sodium cacodylate, 1 mM MgCl2, 1 mM
-mercaptoethanol, pH 7.2) for 15 min at room
temperature. The end-labeling reaction was done by incubating the slides
with the reaction buffer containing 10 µM biotin-dUTP (Boehringer Mannheim GmbH, Mannheim, Germany), and 0.3 U/µl of terminal deoxynucleotidyl transferase (Toyobo Co., Osaka, Japan), for 60 min at 37°C. The slides were washed extensively, blocked with 20% FCS, and then the incorporated biotin was detected using FITC-conjugated streptavidin as in
the protocol for FISH (see below). The slides were treated with RNase A
(100 µg/ml, 37°C, for 20 min), counterstained with PI, and then observed
under the conditions used for FISH.
Probe Preparation and FISH
Preparation of probe from purified micronuclei was as described (Shimizu
et al., 1996), except that DNA in the purified micronuclei was directly
used for biotin labeling by using the BioPrime DNA Labeling System
(Life Technologies Inc., Gaithersburg, MD). C-myc cosmid DNA probe
was prepared as described (Shimizu et al., 1996
). FISH using standard
methanol/acetic acid-fixed nuclei was performed as described previously
(Shimizu et al., 1996
). Assessments of DM localization by confocal microscopy required the following procedure to preserve the spherical shape of
the nuclei. This protocol, based on that developed for human lymphocytes
(Ferguson and Ward, 1992
; Vourc'h et al., 1993
), could not be applied directly to COLO 320DM due to severe nuclear aggregation. The modified procedure involves pelleting 10 ml of COLO 320DM cells by centrifugation at 260 g for 5 min, followed by complete removal of the supernatant. The cells were gently suspended in 50 µl of growth medium and then 10 ml of prewarmed (37°C) 75 mM KCl, 2 mM CaCl2 was added slowly. The
suspension was centrifuged immediately as described above and the supernatant was removed completely. The cell pellet was loosened gently,
suspended in 1 ml of 75 mM KCl and 2 mM CaCl2 at 4°C, and then followed by addition of 1 ml of 75 mM KCl, 2 mM CaCl2, 0.5% Triton X-100
at 4°C. The suspension was kept on ice for 10 min, then Dounce homogenized (loose fitting pestle, 5 times, at 4°C; Fisher, Pittsburgh, PA). 1.5 vol
of 5% PFA in PBS was added to the suspension and then incubated for 10 min at room temperature with occasional gentle shaking. After incubation, a 1:10 vol of 1 M Tris-HCl, pH 7.4, containing 1% BSA was added and further incubated for 10 min at room temperature with gentle shaking. The fixed nuclei were washed twice with PBS containing 1% FCS and
then stored at 4°C up to 1 wk. Before FISH hybridization, the fixed nuclei
were sedimented by cytospin onto poly L-lysine-coated glass slides (Matsunami Glass Ind., Ltd., Osaka, Japan). Slides were treated with RNase A
(Sigma Chemical Co., 100 µg/ml in 2× SSC, 37°C for 60 min), washed
once with 2× SSC for 3 min, and then followed by blocking with 3% BSA
in PBS for 30 min at 37°C. Slides were incubated in 50% formamide dissolved in 2× SSC for 30 min at room temperature to enable buffer equilibration, followed by addition of the hybridization mixture containing labeled probe (prepared as for standard FISH [Shimizu et al., 1996
]). The
sample was covered by a coverslip, sealed completely with rubber cement, denatured at 85°C, and then hybridized using overnight incubation at
37°C. Washing and the detection of the hybridized probe were performed
as described previously (Shimizu et al., 1996
). In some cases, intact cells
were fixed directly and then hybridized. For this purpose, the cells were
cytocentrifuged onto poly L-lysine-coated slides, fixed with ethanol/acetic
acid (19:1) for 3 min at
20°C, rehydrated with PBS, and then treated
with 4% formaldehyde in PBS for 10 min at 4°C. The slides were then
washed extensively with PBS and hybridized as described above for isolated nuclei. Images were obtained using a Bio-Rad MRC600 confocal
system (Hercules, CA) on a Zeiss Axiovert 135 microscope (see Fig. 6).
Most images were obtained using a 63× objective (Apochromat, 1.40, oil,
Carl Zeiss, Inc.), and zoom factor two. The acquired digital images were
expressed as pseudocolors and then merged using Adobe Photoshop (Adobe Systems Inc., Mountain View, CA).
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Localizing DMs in Interphase Nuclei
The locations of DMs in confocal nuclear sections were determined by measuring the distance from each of the hybridized signals to the center of the nucleus using the corresponding nuclear diameter as a unit length. Coplanar PI (DNA) and FITC (hybridized signal) images intercepting the center of the nucleus were obtained from randomly chosen nuclei. The digital images were merged using COMOS software (Bio-Rad Laboratories). The threshold value for each FITC signal was lowered until each signal, representing the domain of one or more DMs, became a single dot to enable accurate distance measurements. The distances from this dot to the center of the nucleus and the nuclear diameter were determined. The location of each signal in the nucleus was expressed by dividing the former number by the latter number. According to this expression, the center of the nucleus is 0, and the outer edge of the nuclear membrane is 1. At the same time, the intensity of each signal was measured using an arbitrary unit scale. These values were determined for every signal in each of the nuclear sections using a minimum of 100 randomly chosen nuclei for each sample. This procedure gives rise to a distribution of DM signals in each two-dimensional focal plane. The heights and widths of each nucleus were found to be approximately equal, indicating that the fixation procedure preserved a spherical nuclear morphology. We assume that the distribution of signals within each nuclear vol should correspond to the number of signals we detected at the corresponding radius in two-dimensional space. Therefore, we corrected the number of signals at each radial location to represent the number that should be present in the spherical volume corresponding to that radius.
Simultaneous Determination of DM Location and DNA Replication Using FISH and BrdU Incorporation
Rapidly growing COLO 320DM cultures were pulse labeled using 10 µM BrdU (Sigma Chemical Co.) for 1 h and then followed by immediate cell collection. The pulse-labeling period was 30 min for the experiment presented in Fig. 2, G and H. The isolation of nuclei, fixation by PFA, and FISH using purified micronuclei probe were done as described above. After FISH, BrdU incorporation was determined by incubating the slides with anti-BrdU mouse monoclonal antibody (PharMingen, San Diego, CA) at a final concentration of 10 µg/ml in PBS containing 0.1% BSA. After a 60-min incubation at 37°C, the slides were washed three times with PBS for 5 min each. The slides were then treated with rhodamine-labeled anti-mouse Ig (Boehringer Mannheim Biochemicals) at a final concentration of 10 µg/ml in PBS containing 0.1% BSA. The slides were incubated for 60 min at 37°C and then washed with PBS three times at 5 min each. The nuclei were viewed without counter staining, using an MRC 1024 (Bio-Rad Laboratories) confocal system equipped to Axiovert 135M microscope (Carl Zeiss, Inc.), and then the acquired digital images were processed as described above.
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Results |
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Nuclear Budding during Interphase Can Selectively Entrap DMs
The classic mechanism by which acentric chromosome
fragments such as DMs are lost from cells involves their
enclosure within reforming nuclear membranes subsequent to telophase (for review see Heddle et al., 1991).
However, there is one report that nuclear anomalies resembling micronuclei can be generated in interphase subsequent to
irradiation (Duncan et al., 1985
). We used a
cell line with amplified c-myc genes to assess the relative
contributions of postmitotic and interphase mechanisms to
DM micronucleation.
A FISH analysis of COLO 320DM, a colon cancer cell
line of neuroendocrine origin, is shown in Fig. 1. A biotinylated FISH probe specific for the c-myc amplicon in
COLO 320 cells was obtained from micronuclei purified
from COLO 320DM cells (Shimizu et al., 1996). FISH analysis with this probe showed that >95% of the cells in the
population contained only DMs, and the remainder contained DMs along with one intrachromosomally amplified
region (Fig. 1 A, arrow). Consistent with a previous report
(Levan and Levan, 1978
), the DMs in the prometaphase
spread (Fig. 1 A) do not appear to be distributed randomly
since many localize to the periphery of the prometaphase
ring. Peripheral nuclear localization was also observed in
interphase nuclei using confocal microscopy (see below).
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Analyses of exponentially growing cultures of COLO
320DM cells by a conventional FISH procedure revealed
that 2.8 ± 0.9% of interphase nuclei have projections or
buds. These buds can be classified into two types by FISH
analysis using the probe described above. The first class
(56 ± 12.2% of the nuclear buds) contained highly concentrated DM sequences (Fig. 1, B-D show representative buds that contain DMs; C shows two micronuclei and one
bud). We also observed a second class of buds that were
stained with PI, but did not hybridize with the DM probe.
This suggests that DNA that does not hybridize with the
DM probe was contained in these buds. (see Discussion).
However, the selectivity for acentromeric sequences such
as DMs appears to be very high. This is indicated by the
nuclear bud shown in Fig. 1 D that contains three DNA
clusters that stain with PI (the red signal represents DNA
staining as these samples were first treated extensively
with RNAse) and that hybridize intensely with the micronuclear DNA-FISH probe (Fig 1 D'; merge shown in
D''). The selective inclusion of DMs into buds was also
readily apparent when we used a FISH probe derived from a cosmid containing the c-myc gene. (Fig. 1 E), PFA-fixed nuclei isolated by a hypotonic method (see Fig. 6), or
an isotonic method (data not shown). Furthermore, the selective inclusion of DM sequences into nuclear buds was
also apparent in intact cells fixed directly by ethanol/acetic
acid (19:1) followed by formaldehyde (Fig. 1, F and G).
The apparent preferential inclusion of DM sequences in
buds is reminiscent of our previous finding that DMs are
highly enriched in micronuclei and that chromosomes or
minichromosomes with functional centromeres are excluded from micronuclei (Von Hoff et al., 1992; Shimizu et
al., 1996
). These data suggest that buds are precursors of
micronuclei.
Nuclear Buds and Micronuclei Are Formed during S Phase
The nuclei with buds exhibit a morphology typical of an
interphase cell, not one in mitosis. Therefore, we determined the kinetics of formation of micronuclei and nuclear buds to ascertain whether these structures can be
generated during S phase. COLO 320DM cells were synchronized at the G1/S boundary using a two-step procedure involving treatment with high thymidine concentration to arrest cells during S phase, release for 12 h to
enable progression through and exit from S phase, and
then incubation with the DNA polymerase inhibitor
aphidicolin to arrest cells at the beginning of the next S
phase (Stein et al., 1994). Removal of aphidicolin resulted
in rapid entry into S phase with a peak of [3H]thymidine
uptake at 4 and a peak of mitosis at 10 h, respectively (Fig.
2 A). The cells entered a second, less synchronous cycle
19 h after release. The synchronization level of the first S phase was further quantified by BrdU-pulse labeling followed by confocal microscopic examination of the nuclear
labeling patterns. The labeling pattern progressed as reported previously (O'Keefe et al., 1992
), with six readily
distinguishable patterns (Fig. 2 G, Patterns 0-6). Examination of the arrested cells (t = 0) revealed that 96.1% of the
cells did not incorporate label (Fig. 2 H, Pattern 0) but the
remaining 3.9% of cells exhibited the pattern expected for
early S phase (Fig. 2 G, Pattern 1). The small but significant fraction of cells in early S phase most likely reflects
the leakiness of the synchronization procedure. After the
release from the aphidicolin block, these patterns progressed sequentially as shown in Fig. 2, G and H. At the
peak of [3H]thymidine uptake (4 h after release), 8.3% of
cells still did not incorporate BrdU, suggesting that some
cells may have been arrested irreversibly by aphidicolin.
The frequencies of micronuclei and buds were ascertained using the DNA specific dye DAPI (Fig. 2, C and D). We determined whether these structures contain amplified sequences by hybridizing all samples with the DM-painting probe obtained from micronuclei. The frequency of nuclei with buds at the G1/S boundary (i.e., t = 0) was nearly zero, increased dramatically as the cells progressed through early S phase (t = 0-5 h), declined in later S phase, and then gradually increased as the cells entered and progressed through a second S phase (Fig. 2 C). FISH analysis demonstrated that the buds enriched in DM sequences (DM + buds) showed the same time course (Fig. 2 E). The frequency of buds peaked concurrent with a BrdU labeling pattern characteristic of the onset of replication of peripheral heterochromatin (Fig. 2, G and H, Patterns 3). Importantly, the number of total micronuclei, or those with DMs, increased and declined in register with the number of nuclear buds (Fig. 2 C), suggesting that buds are precursors of micronuclei. We recently found that micronuclei containing DM sequences are released into the growth medium. The extracellular micronuclei were distinct from apoptotic bodies since they did not have condensed chromatin, the DNA was not degraded extensively, and the nuclear lamina were still intact (Shimizu, N., and G.M. Wahl, manuscript in preparation). Therefore, it is reasonable to propose that the decrease of micronuclei in late S phase reflects their release from the cells.
Whereas the frequency of nuclei with buds was low at
the G1/S boundary, micronuclei were readily apparent
(Fig. 2 C). One likely possibility is that these micronuclei
were generated during the telophase of a previous cell cycle. We modified the synchronization strategy to include
nocodazole to block cells in prometaphase after release
from the aphidicolin block. This protocol restricts the
analysis to micronuclei and buds generated within a single S phase, and to prevent buds or micronuclei produced
from cells arrested before prometaphase of the previous
cycle from entering the analysis (Cassimeris and Salmon,
1991). The treated cells entered and progressed through S
phase at approximately the same rate as those not exposed
to nocodazole (Fig. 2, compare A and B), and the mitotic
index increased significantly by 8 h after release. The effectiveness of nocodazole treatment is also indicated by
the inability of the drug-treated cells to progress into a second S phase over the time course used. Importantly, the
number of nuclei with buds was again nearly zero at the
G1/S boundary, and both the number of buds and micronuclei increased after release into nocodazole-containing
medium. These data are consistent with the interpretation
that nuclear buds and micronuclei can arise de novo during S-phase progression.
The kinetic and nuclear morphologic analyses of budding and micronucleation indicate that both events can occur during S phase. Since slowing replication fork progression may lead to DNA breakage (Eki et al., 1987; Linke et
al., 1996
), and micronuclei preferentially capture acentric
fragments (Von Hoff et al., 1992
; Shimizu et al., 1996
), we
determined whether replication inhibitors increase S phase
micronucleation efficiency. The drugs tested included inhibitors of ribonucleotide reductase (HU, deferoxamine,
and guanazole), an inhibitor (PALA) of the carbamyl
phosphate synthatase, dihydro-orotase, aspartate transcarbamylase (CAD) enzyme complex that catalyzes the first
three steps of de novo pyrimidine biosynthesis, and aphidicolin. DNA synthesis inhibitors produced substantial increases in micronucleation, and this generally correlated
with an increased fraction of cells in S phase (Fig. 3, A and
B). A sharp decrease in micronucleation efficiency was observed for deferoxamine and PALA when these drugs
were used at concentrations that severely inhibited S
phase (Fig. 3 B and data not shown). These data indicate
that micronucleation can result from inhibitors that retard
replication fork progression and lengthen S phase.
We analyzed the effects of inhibitors that do not affect
DNA synthesis to ascertain whether micronucleation can
result from interfering with other DNA transactions such
as DNA repair, or by interfering with membrane structure. Two inhibitors of poly(ADP-ribose) polymerase
(nicotinamide and coumarin) were tested since ADP ribosylation has been implicated in DNA repair (Satoh and
Lindahl, 1992), and inhibiting repair could increase the
probability of generating acentric chromosome fragments.
We tested the effects of DMSO, a membrane-active polar
compound previously reported to reduce DM copy number in some tumor cell lines (Shima et al., 1989
; Eckhardt et al., 1994
). DMSO increased micronucleation without
lengthening S phase (Fig. 3, A and B). Coumarin had no
effect on micronucleation, whereas nicotinamide produced a small increase under conditions that apparently
increased the amount of damage in the cells since there
was a significant increase in the G2 fraction (Fig. 3, A and
B). These data are consistent with the view that micronucleation efficiency can be increased by at least two mechanisms, one of which presumably involves perturbing replication fork progression, and another of which may involve
events occurring outside of S phase.
Peripheral Nuclear Localization of DMs Correlates with Their Elimination by Budding
Insight into the mechanisms underlying the selective inclusion of DMs into nuclear buds and the formation of these
structures in interphase was obtained by confocal microscopy. PFA fixation of nuclei was used for optimal preservation of nuclear morphology (Manuelidis and Borden,
1988).
Confocal sections from three representative nuclei isolated from rapidly growing untreated COLO 320DM cells
are shown in Fig. 4, A-C. FISH revealed preferential localization of most DM sequences to the nuclear periphery, as
indicated by the significant hybridization intensity, clustering, and number of DM signals at the extreme edge of
each nucleus. Note the substantial deviation from a random distribution of DM sequences at the nuclear periphery (quantified in Fig. 5 A by measuring DM positions relative to the center of each nucleus in 100 interphase
nuclei). By contrast, sequences amplified within chromosomes in the closely related cell line COLO 320HSR
showed a nearly random distribution throughout the nucleus (Fig. 5 C). HU treatment preferentially depleted
DMs from the nuclear periphery (Fig. 4, D-F and Fig. 5 B,
quantification) and then reduced the DM content per cell
by approximately threefold as determined by competitive
PCR amplification (for method see Shimizu et al., 1996;
data not shown). Taken together with the data reported
above, these results indicate that DM sequences located at
the nuclear periphery are preferentially incorporated into
nuclear buds, which are then removed from the nucleus
through the formation of micronuclei.
|
Incorporation of Replicating DM Sequences into Nuclear Buds
The correlations between S phase progression, nuclear
budding, and micronucleation reported above led us to investigate whether DM sequences undergoing replication
are targeted for inclusion into buds. This possibility was
examined by pulse labeling COLO 320DM cells with
BrdU, and then hybridizing the isolated nuclei with the
DM-FISH probe. Subsequent reaction with an anti-BrdU antibody and fluorescein-labeled secondary antibody enabled simultaneous detection of nuclei, buds, and micronuclei containing DMs that were undergoing DNA replication during the brief labeling interval. FISH analysis of
two representative confocal sections (Fig. 6, A-A'' and
B-B'') shows that nuclear buds in these cells (arrows) contain highly concentrated DM sequences. The nuclei, nuclear buds, and peripheral regions of each nucleus incorporated
BrdU, indicating that these buds were formed in nuclei
that were synthesizing DNA at the time of bud formation.
Table I shows that BrdU+, DM+ buds (type 1, 1') represent 48% of the total population of DM-containing buds.
Some nuclei incorporated BrdU, but the buds they produced were not labeled (type 2, 2'; 35%). We infer that
some of these buds were also generated during S phase (as
opposed to during the previous cycle), and that they did
not reveal BrdU incorporation because their DNA was
not undergoing replication during the brief BrdU incubation period. Samples in which neither nuclei nor buds labeled with BrdU may represent examples where buds
were generated outside of S phase (type 3; 17%). Alternatively, the presence of such buds (type 3) may indicate that
these structures persist after DNA replication has finished,
and may be sources of some of the micronuclei observed
to form during mitosis. These data reveal a strong correlation between DMs undergoing replication and their inclusion in buds and micronuclei, and they lead to a conservative estimate of 50% of the micronuclei produced during each cell cycle being generated during S phase.
|
The S phase micronucleation process described here superficially resembles the induction of "nuclear anomalies"
by an apoptotic mechanism following irradiation (Duncan et al., 1985
) or colchicine treatment (Duncan et al.,
1984). However, the absence of highly condensed DNA in
the nuclei-producing buds suggests that they were not undergoing apoptosis at the time of bud formation. To determine whether budding and micronucleation are separable
from apoptosis, we determined whether buds contain the
condensed, fragmented DNA that typifies apoptotic cells.
Fragmented DNA was detected using the TUNEL assay
in which terminal transferase is used to add BrdU to the
3'-OH groups generated by apoptotic DNA fragmentation (Gavrieli et al., 1992
). The cells were also stained with PI
to visualize all nuclei and buds. An example of the data
obtained from such an analysis of COLO 320DM cells is
shown in Fig. 6, C-C''. The TUNEL assay shows one cell
with a lobular nucleus that exhibits a strong TUNEL reaction and typifies the fragmented, condensed DNA observed in apoptotic nuclei (Cohen, 1993
). The PI staining
in the middle panel reveals a cell producing a nuclear bud
that does not stain by the TUNEL assay and does not exhibit the pycnotic structure of the apoptotic nucleus. Synchronization experiments showed that whereas ~5% of
cells generated buds at the peak of S phase (e.g., refer to
Fig. 2 C), only 0.5-1% were TUNEL positive. Other data
(see Discussion) provide additional evidence that S phase
budding and micronucleation do not require activation of
an apoptotic program.
Micronucleation Occurs Infrequently in Normal Cells, and Is Increased upon p53 Inactivation
Micronucleation occurs at significantly lower rates in normal cells than tumor cell lines (Roser et al., 1989; Bondy et
al., 1993
). Since micronucleation can be induced by chromosome breakage (Heddle and Carrano, 1977
), the observed increase in micronucleation in tumor cells might result from mutations that increase the probability of DNA
breakage. The tumor suppressor p53 controls G1 arrest responses activated by DNA breakage and rNTP depletion
induced by PALA treatment, and DNA breakage can occur in p53-deficient cell lines that enter S phase during
PALA treatment (Livingstone et al., 1992
; Yin et al., 1992
;
Linke et al., 1996
). As reported above, PALA also induces S-phase micronucleation in COLO 320 cells. These data
led us to investigate whether p53 inactivation in normal
diploid fibroblasts results in increased S-phase budding
and micronucleation.
Human WS1 normal diploid fibroblasts, and two nearly
isogenic derivatives generated by retroviral transduction
of the neomycin phosphotransferase gene (WS1-neo) or
an oncogenic derivative of the human papilloma virus E6
gene (WS1-E6) were used. The E6 gene product facilitates
p53 degradation by a ubiquitin dependent pathway (Scheffner et al., 1990; Crook et al., 1991
). Cell cycle checkpoint controls that regulate entry into S phase in the presence of
DNA damage or limiting rNTP concentrations appear to
be inactivated to equivalent degrees in human cells expressing mutant p53, oncogenic E6 protein, and mouse
embryo fibroblasts with homozygous p53 knock out (Kastan
et al., 1992
; Kuerbitz et al., 1992
; Livingstone et al., 1992
;
Yin et al., 1992
; White et al., 1994
; Linke et al., 1996
; Linke
et al., 1997
). Importantly, we previously showed that the
frequency of
radiation induced micronucleation is higher in p53
/
MEFs than in wild-type MEFs (Huang et al.,
1996
). It is likely, therefore, that effects on micronucleation observed upon expression of oncogenic E6 protein
relate to inactivation of p53 rather than to other proteins
that may be affected by E6.
The data shown in Fig. 7 demonstrate that E6 gene expression increases the micronucleation rate of WS1 cells.
WS1 cells exhibit a low micronucleation rate that is not increased by HU or PALA (Fig. 7 A). Consistent with our
previous studies (Linke et al., 1996), PALA induced a G1
cell cycle arrest, whereas HU did not significantly affect
the percentage of WS1 cells in S phase at the concentration used (Fig. 7 B). By contrast, E6 expression increased
the micronucleation efficiency of these fibroblasts growing
under normal conditions, and both HU and PALA produced a substantial further increase in micronucleation
rate (Fig. 7 A), which correlated with a significant increase
in the number of cells in S phase (Fig. 7 B). The importance of an E6 target, which we infer to be p53, in limiting
micronucleation is evident in other cell types since similar
results were obtained using RPE-h and their E6 expressing derivatives (data not shown).
|
The elongation of S phase and induction of micronuclei
by HU and PALA in both COLO 320DM and WS1-E6
cells led us to assess whether budding in S phase is the predominant mechanism of micronucleation in WS1-E6 cells.
WS1-neo and WS1-E6 cells were arrested in G0 by serum
deprivation and then released in the presence of aphidicolin to arrest the cells at the G1/S boundary (Fig. 7 C).
Synchronization by serum depletion did not increase micronucleation rate (data not shown) and then the micronucleation and budding frequencies did not increase in S
phase in WS1-neo cells (Fig. 7 D). By contrast, removal of
aphidicolin from WS1-E6 cells resulted in significant increases in the frequencies of both nuclear budding and micronucleation as the cells progressed through S phase (Fig.
7 D). Since DNA damage does not induce apoptosis in either WS1 or WS1-E6 cells (Di Leonardo et al., 1994; Linke
et al., 1996
; Linke et al., 1997
), the increased S-phase micronucleation observed in these cells occurs independent
of an apoptotic program.
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Discussion |
---|
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---|
Loss of cell cycle checkpoints during cancer progression
creates a permissive environment for the initiation and
propagation of chromosomal rearrangements such as amplification of cellular protooncogenes. The persistence or
elimination of these structures within the nucleus can, respectively, promote or inhibit cancer progression. Interestingly, in human tumors analyzed at biopsy, oncogene amplification occurs most frequently in acentric chromosomal fragments such as DMs (Benner et al., 1991), and micronucleation represents a major pathway for the elimination of
such structures (Von Hoff et al., 1992
; Shimizu et al., 1996
).
Before this report, micronucleation had been considered
to result from imperfect segregation of acentric chromosomal fragments or fragments of overly long chromosomes
during karyokinesis (Heddle and Carrano, 1977
; Heddle et
al., 1983
). The results presented here, by contrast, reveal that acentric DMs are sorted to the nuclear periphery during S phase, and are then selectively eliminated from the
nucleus by micronucleation in advance of karyokinesis.
S Phase Budding and Micronucleation May Be the Predominant Mechanism for DM Elimination
We infer that S-phase micronucleation is a general characteristic of human cells with an aberrant p53 pathway since
this process was observed in a human tumor cell line, and
p53-deficient normal human fibroblasts and epithelial
cells. The cell synchrony and release experiments provide
direct evidence that budding begins as cells enter S phase.
The experiment summarized in Table I shows that BrdU+
micronuclei account for almost 50% of the total micronuclei generated during a single S-phase. It is reasonable to
infer that a fraction of the 35% of BrdU micronuclei derived from BrdU+ nuclei did not incorporate the label because their DNA did not replicate during the brief BrdU
pulse used. Such micronuclei would, therefore, also have
originated during S phase, but we have no means of providing a precise estimate of this fraction. The available
data do, however, demonstrate that S-phase micronucleation mediated by budding is at least as common as the
classic postmitotic process, and may be the predominant
mechanism for removing DMs from human cancer cells.
The parallel increases in budding and micronucleation frequencies as cells enter and proceed through S phase, and the absence of a significant temporal lag between the two, is consistent with a precursor-product relationship in which a micronucleus is produced shortly after a bud is generated. The decrease in budding and micronucleation during the latter part of S phase is also consistent with a tight linkage to the replication program, and suggests that once micronuclei are generated during S phase, some may be released from the cell, whereas others may fuse to nuclei or be degraded intracellularly. Among these possibilities, we have direct experimental evidence that some micronuclei may be expelled from the cell through the cytoplasmic membrane since we have observed extracellular micronuclei containing three membranes and amplified c-myc genes (Fig. 8; Shimizu, N., and G.M. Wahl, unpublished data). We are currently testing whether such micronuclei are bound by a cytoplasmic membrane, and whether they can fuse to the same or other cells in the population (Fig. 8).
|
The baseline frequency of micronuclei observed in cultures at the beginning of S phase in which budding was not
yet evident suggests that some micronuclei were generated
in a previous cycle and persisted for an extended time,
which would be consistent with previous reports of long
lived micronuclei (Heddle et al., 1983). It is conceivable,
therefore, that there are two classes of micronuclei that
differ from each other by their stability, perhaps because
of structural distinctions deriving from their mechanisms
of formation. Consistent with the inference of multiple types of micronuclei, we found that micronuclei can differ
in their lamin and nuclear pore contents (Shimizu, N., and
G. Wahl, unpublished observations). Experiments in progress
are designed to elucidate whether such differences correlate with stability and alternative mechanisms of formation.
S-phase Budding and Micronucleation Do Not Require Previous Engagement of an Apoptotic Program, but Can Result in Apoptosis
Apoptosis can generate nuclear blebs (Dini et al., 1996)
and has been inferred to produce "nuclear anomalies" that
resemble micronuclei (Duncan and Heddle, 1984
; Duncan
et al., 1985
). However, our analyses show that nuclei that
produced buds were not pycnotic and fragmented like
apoptotic nuclei. Buds and micronuclei generated in
COLO 320DM cells within a single S phase were not
TUNEL positive, indicating that they did not contain fragmented DNA. Whereas apoptotic cells did arise in HU-treated COLO 320DM cultures, this required prolonged
incubation, and occurred after a substantial fraction of the
amplified c-myc genes had been removed. Furthermore,
cells undergoing budding and micronucleation survived
for several days, which is not expected if an apoptotic program were activated before micronucleation. A time
course experiment similar to that described above (e.g.,
Fig. 2) revealed that budding increases during a single S
phase whereas the fraction of cells undergoing apoptosis remained roughly constant, and that most of those generating buds were not TUNEL positive (data not shown). Finally, though we have not observed apoptosis in normal fibroblasts (Di Leonardo et al., 1994
), and loss of p53
function typically makes cells more resistant to apoptosis
induced by growth conditions that can lead to DNA damage (White, 1994
), S-phase budding and micronucleation
were induced in normal diploid fibroblasts upon expression of oncogenic papillomavirus E6 protein, presumably
due to elimination of p53 function. These observations
lead us to propose that buds and micronuclei in COLO
320DM and WS1-E6 cells are produced by a mechanism that does not require prior engagement of the apoptotic
program.
Mechanisms of Budding and Micronucleation
The molecular basis for inclusion of DMs in buds and micronuclei, and exclusion of similarly sized centric fragments from such structures (Shimizu et al., 1996), remains
to be elucidated. We offer two potential explanations. The
first relates to replication of DMs at an inappropriate nuclear location. Previous studies showed that heterochromatic DNA, such as that comprising inactive X chromosome-Barr bodies, typically replicates late in S phase at
the nuclear periphery and is often enclosed within micronuclei, whereas the active X replicates earlier in S phase at
a more internal position and is not subject to micronucleation (Dyer et al., 1989
; Tucker et al., 1996
). DMs are euchromatic and tend to replicate early in S phase, typically
at approximately the same time as the native chromosomal locus (Carroll et al., 1991
, 1993
). However, as shown
here, they can localize to and replicate at the nuclear periphery. Since chromosomes occupy specific territories
within interphase nuclei (Cremer et al., 1993
), the independence of DMs from chromosomes may prevent them
from occupying the correct nuclear position and could explain their localization to the periphery. Peripheral localization may be a default position reflecting the inability of
DMs to undergo the nuclear movements choreographed by centromeres and/or telomeres during S phase (DeBoni
and Mintz, 1986
; Ferguson and Ward, 1992
; Vourc'h et al.,
1993
). Replication at such peripheral locations, or perhaps
inability to move away from such sites into protected inner
nuclear regions following replication, may then precipitate
bud formation. A second explanation is that some chromosomal sequences may bind proteins that target the protein and associated nucleic acid to a particular nuclear location, such as the periphery. In the absence of a centromere/ telomere, this protein-nucleic acid complex may be destined to form a nuclear bud until membrane fusion produces a micronucleus. This view is supported by recent
studies in Tetrahymena demonstrating that cis-acting heterochromatic regions bound by the chromodomain protein Pdd1p are targeted to the nuclear periphery when the
DNA is fragmented during macronuclear development
(Madireddi et al., 1996
). The peripheral, Pdd1p-associated
acentric chromosomal fragments are then removed from
the nucleus via micronucleation. The striking similarities
between the observations in Tetrahymena and those reported here raise the intriguing possibility of an evolutionarity conserved process that distinguishes intact chromosomes from chromosome fragments or other acentric
DNA such as DNA viruses to facilitate removal of the latter from the nucleus.
S-Phase Micronucleation Suggests That Loss of p53 Function Affects the Probability of Chromosome Breakage
Treatment of normal fibroblasts with HU or PALA did
not induce S-phase micronucleation, whereas identical
treatment of isogenic p53-deficient normal fibroblasts increased S-phase micronucleation frequency significantly.
These data, along with previous observations, lead us to
propose that this micronucleation increase most likely reflects a higher probability of DNA breakage occurring in
the p53-deficient cells when they attempt DNA replication
under adverse conditions. In support of this idea, micronucleation is an indicator of chromosome breakage, and has
long been used as an assay for clastogens (Heddle et al.,
1991). Inhibition of replication fork progression induces
chromosome breakage in bacteria, yeast, and mammals
(Eki et al., 1987
; Kuzminov, 1995
; Michel et al., 1997
). Furthermore, replication inhibitors including PALA, methotrexate, and aphidicolin can induce chromosome breakage
through expression of fragile sites (Kuo et al., 1994
; Coquelle et al., 1997
). Although the relationship between
fragile site induction and p53 function has not been tested,
the cell lines used for such studies were competent for
gene amplification and consequently should have had a
defective p53 pathway (Livingstone et al., 1992
; Yin et al.,
1992
). Consistent with a breakage-moderating function of p53, we previously reported that PALA generates chromosome damage in cells with defective, but not normal,
p53 function (Linke et al., 1996
). Taken together, the data
lead us to propose that p53 minimizes the frequency at
which structural chromosomal alterations are induced during exposure to suboptimal growth conditions by at least
two mechanisms. First, as shown previously, p53 mediates a G1 arrest in response to low rNTP pools, which prevents
cells from entering S phase with inadequate precursors for
DNA replication (Linke et al., 1996
). Second, the data
presented here suggest that p53 minimizes S-phase DNA
breakage in cells that attempt DNA replication during diverse metabolic challenges. The mechanisms underlying
the proposed S-phase function of p53 are currently under
investigation.
Micronucleation and Cancer Treatment
Drugs that enhance S-phase budding should prove valuable for chemotherapy as they affect a process (micronucleation) and cytogenetic aberration (DMs) that are restricted to cancer cells. However, their efficacy is limited
by the extent to which DM removal prevents further cell
growth, or enhances sensitivity to other therapeutic strategies. In COLO 320DM cells, induction of the S-phase budding mechanism can decrease the number of DMs sufficiently
to reduce plating efficiency in soft agar and tumorigenicity
in vivo (Von Hoff et al., 1992). Interestingly, COLO 320HSR
cells, which have approximately the same number of c-myc
genes amplified within chromosomes, did not exhibit such
phenotypic changes upon HU treatment and did not decrease c-myc copy number, even though they produced approximately the same number of micronuclei as COLO
320DM cells (Von Hoff et al., 1992
). The same HU treatment conditions also induced apoptosis more rapidly and
in a higher fraction of COLO 320DM than COLO 320HSR
cells (data not shown). Similarly, HU treatment of HL-60
DM, but not HL-60 HSR cells, reduced c-myc copy number, induced differentiation, and later, apoptosis (Eckhardt et al., 1994
; Shimizu et al., 1994
). These results suggest that reduced tumorigenicity may result from induction
of an apoptotic program when a sufficient number of extrachromosomally amplified sequences encoding oncogenes are removed from the cell. The recognition of a
mechanism for the segregation and elimination of amplified sequences and other acentric DNA macromolecules,
and the recognition that some agents can enhance this process, present important opportunities to expand and refine
chemotherapeutic strategies, and to gain insight into the
cis-acting elements and trans-acting factors that determine
nuclear DNA localization.
![]() |
Footnotes |
---|
Received for publication 7 October 1997 and in revised form 16 December 1997.
We would like to acknowledge S.P. Linke (National Cancer Institute, Bethesda, MD) for kindly providing cell lines, D. Peterson (Salk Institute, La Jolla, CA) for his kind help on the operation of confocal microscopy that appeared in Fig. 6, and D. Von Hoff and K. Davidson (both from Institute of Drug Development, University of Texas Health Science Center, San Antonio, TX) for allowing us to cite their unpublished results. We thank T. Shimura and N. Kumon (both from Hiroshima University, Higashi-Hiroshima, Japan) for their technical help. T. Paulson, T. Kanda, S. O'Gorman, S. Pfaff, G. Karpen, O. Vafa, and L.-c. Huang (all from Salk Institute except Paulson [Fred Hutchinson Cancer Research Center, Seattle, WA] and Huang [UBI, Menlo Park, CA]) provided helpful discussions concerning experimental procedures and topics presented in this manuscript. ![]() |
Abbreviations used in this paper |
---|
BrdU, 5-bromo-2'-deoxyuridine; DAPI, 4'-6-diamidino-2-phenylindole; DM, double-minute chromosome(s); FISH, fluorescence in situ hybridization; HU, hydroxyurea; PALA, N-phosphoracetyl-L-aspartate; PFA, paraformaldehyde; PI, propidium iodide; RPE-h, normal human retinal pigmented epithelial cells.
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