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Carcinogenesis |
Departments of Clinical Pharmacology [H. Z., A. R. B.] and Molecular Toxicology [N. J. G.], Imperial College School of Medicine, London, SW7 2AZ United Kingdom
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ABSTRACT |
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INTRODUCTION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Like all other HCAs, PhIP requires metabolic activation to become mutagenic. The major pathway for the activation involves N-hydroxylation catalyzed by cytochrome P450, followed by esterification of the N-hydroxy intermediate (15, 16, 17, 18) . The activated PhIP attacks and covalently binds DNA, primarily at the C-8 position of guanine forming the N2-(deoxyguanosin-8-yl)-PhIP (19, 20, 21, 22) . The formation of such DNA adducts is thought to be premutagenic.
DNA damage can elicit a range of cellular responses, one of which is cell cycle arrest. Such a cellular response is considered to be an essential defense mechanism. Key cell-cycle-dependent proteins maintain checkpoints that allow cells to repair damaged DNA or allow them to die by apoptosis if the damage is too severe. It has been proposed that p53/p21WAF1/CIP1 is involved in G1-S checkpoint control (23) and E2F/cyclin A-cdk2 is involved in S-G2 checkpoint (24) . The misfunction or loss of these control points can result in inefficient or ineffective DNA repair leading to genomic instability, the inheritance of genetic change, and the development and progression of neoplastic disease. HCAs such as PhIP are clearly potent chemical carcinogens, capable of damaging DNA resulting in mutation, but whether they can also interfere with the maintenance of cellular responses to DNA damage is not clear. Therefore, to better understand the processes through which the HCAs exert their carcinogenic potential, we have examined the early cellular events and gene mutations elicited in human lymphoblastoid cells by exposure to metabolically activated PhIP.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cell Lines.
TK6 human lymphoblastoid cells were obtained from the European
Collection of Cell Cultures (Wiltshire, United Kingdom) and maintained
in suspension at a density of 0.5–1 x 106/ml in the culture medium (RPMI 1640
supplemented with 10% fetal bovine serum) and 2 mM
glutamine, Life Technologies, Inc., Paisley, United Kingdom). The cells
were diluted daily. XEMh1A2-MZ cell line, a variant of Chinese hamster
fibroblast V79 genetically engineered to express human CYP1A2, was
generously provided by Dr. J. Doehmer (Institut Fur Toxikologie und
Umwelthygiene, Technische Universita, Munich, Germany). Adherent
XEMh1A2-MZ cells were maintained in the same medium supplemented with
0.4 mg/ml of geneticin to select for G418 resistance. Both cell lines
were cultured at 37°C in 5%CO2/95% air.
Treatment.
XEMh1A2-MZ cells were seeded into 6-well plates at 2.5 x 106/well and allowed to attach to the
plate. After 24 h, they were irradiated (50 Gy), and the
supernatants were aspirated just before adding TK6 cells at
2.5 x 106 cells/well in 5 ml of
culture medium. In preliminary experiments, we showed that irradiated
XEMh1A2-MZ cells were able to activate PhIP to N-hydroxy
PhIP and remained adhered to the plate for at least 3- 4 days
with minimal cell loss as evidence by floating cells (determined by
microscopy). Under the conditions of coculture, the XEMh1A2-MZ cells
remained attached and viable with functional CYP1A2 activity but were
unable to replicate, whereas the nonadherent TK6 cells were viable and
grew exponentially. The coculture was treated for 20–40 h with PhIP
(Toronto Research Chemicals Inc., Toronto, Canada; final concentration
0–10 µg/ml, dissolved in DMSO). Nonadherent cells were harvested by
centrifugation, and the cell pellets were collected for further culture
or for cell cycle analysis.
Flow Cytometry Analysis of Cell Cycle Distribution and Cyclin A
Expression.
Cell cycle stage was determined using flow cytometry. Briefly, TK6
cells were fixed with 70% ethanol at -20°C for 1 h. Cells were
resuspended in 1 ml of PBS containing propidium iodide (5 µg/ml) and
RNase A (0.1 mg/ml) and were incubated at 37°C for 30 min. Cells
(104 cells per analysis) were examined by flow
cytometry (Coulter, Hialeah, FL), and the cell cycle distribution was
determined by DNA content. Cells distributed in
sub-G1 were defined as apoptotic according to the
criteria described by others (25)
.
Simultaneous cell cycle and cyclin A expression was determined by flow cytometry (26 , 27) . Briefly, cells (106) were fixed in a 1:1 (v/v) solution of acetone in absolute ethanol for 30 min at -20°C. After washing with 70% ethanol and PBS, the cells were incubated overnight at 4°C with rabbit polyclonal antibody to human cyclin A (Santa Cruz Biotechnology, Inc., California, USA), which was diluted 1:1000 with PBS containing 1% BSA (BSA). The cells were washed with PBS and incubated with FITC (FITC)-conjugated goat antirabbit IgG antibody which was diluted 1:200 in PBS containing 1% BSA, for 1 h at room temperature in the dark. Cells were washed and resuspended in 1 ml of PBS containing propidium iodide (5 µg/ml) and RNase A (0.1 mg/ml), incubated at 37°C for 30 min. The negative control was processed in a similar way, except that rabbit IgG was used instead of the cyclin A antibody. FITC and propidium iodide fluorescence were measured in the same cells and cyclin A-expressing cells were defined using a gated window. Cyclin A expression for the gated population was quantified as the mean labeling fluorescence.
Cell Cycle Synchronization.
TK6 cells were synchronized in G1 by starvation.
Cells were grown in the culture medium for more than 3 days without
change of medium, then released to fresh medium for 4 h prior to
the treatment, when more than 80% of the cells were in
G1, as determined by flow cytometry.
Cell Survival (Colony-forming Ability) Assay.
Cell survival and viability were assessed by colony formation assay in
the culture medium. TK6 cells treated with PhIP or vehicle control were
seeded at 2–20 cells/well in 96-well plates (round-bottomed) in the
presence of 104 cells/well of irradiated (50 Gy)
TK6 cells. Preliminary experiments had shown that the inclusion of
irradiated TK6 cells aided the growth of mutant colonies. Under these
conditions, irradiated TK6 cells failed to proliferate whereas viable
nonirradiated TK6 cells formed colonies that were counted on day 14 or
day 21 to determine the cloning efficiency.
Cloning of 6-TG-resistant Mutants.
Resistance to the lethal effects of the purine analogue 6-TG was used
as the genetic marker for measurement of mutant frequency and selection
of hprt-mutant clones. The assay detects DNA damage that
results in functional changes to the hprt enzyme, including point
mutations, frameshifts, and deletions (28)
. After
treatment with PhIP, TK6 cells were maintained in exponential growth
for 6 days in fresh medium to allow phenotypic expression. To select
for hprt mutations, cells were seeded into selective medium
(culture medium containing 0.8 µg/ml of 6-TG) at
104cells/well in the presence of
104 cells/well of irradiated TK6 cells. In
preliminary experiments, we found that the inclusion of irradiated TK6
cells in the selection plates aided the expansion of mutant colonies.
The 6-TG-resistant colonies were counted at 14 days or at 21 days after
plating to determine cloning efficiency. The mutation frequency was
determined as the ratio of cloning efficiency in selective medium
compared with nonselective medium.
Statistics.
Statistical analysis was performed using ANOVA.
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RESULTS |
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Early Cellular Response to PhIP Treatment: S-Phase Arrest and
Apoptotic Cell Death.
Single-parameter flow cytometry histograms were used to display cell
cycle distribution. Fig. 1
shows representative cell cycle profiles of asynchronized TK6 cells
before and after treatment with DMSO or a range of concentrations of
PhIP for 20 and 40 h. The experiment has been repeated on
more than three separate occasions with similar results. Table 1
describes the same data shown in Fig. 1
, expressed as % cells in each
phase. The time of growth is different for each column of figures but
is identical within a column; therefore, comparisons only within a
column are appropriate. After 20 h of treatment with PhIP
(1.25–10 µg/ml) there was a slight S-phase shoulder on the
G2-M peak (Fig. 1, b-e,
arrows), indicative of S-phase delay. When PhIP treatment
was extended to 40 h, a distinct S-phase peak was visible,
particularly in cells treated with the highest concentration of PhIP
(10 µg/ml; Fig. 1j).
After treatment withdrawal for 5
days, cells that were treated with PhIP for 20 h reached the same
growth status as DMSO-treated samples regardless of the concentration
of PhIP, indicated by the similar cell-cycle profile between DMSO- and
PhIP-treated cells [Fig. 1
(compare p with
l-o)]. In contrast, the abnormal cell cycle induced by
40 h of PhIP treatment continued to deteriorate over the next 4
days despite culture in PhIP-free medium. A mid-S-phase peak was
clearly seen in cells treated with the lower concentrations of PhIP
(1.25–2.5 µg/ml; Fig. 1, q and r).
However, at
the highest concentrations of PhIP (5–10 µg/ml), the cells showed
evidence of severe damage to the cell cycle and the S-phase arrest
became increasingly difficult to visualize (Fig. 1, s and t).
The decrease in the G2-M peak
induced by 40 h of PhIP treatment was concentration-dependent
(Fig. 1, q-t)
and confirmed a blockage from S phase to
G2-M.
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7%) was observed
(Fig. 1, b-e and g-j,
and Table 1
). It has been
proposed that the presence of a sub-G1 cell population is indicative of
apoptotic cells (25)
. Flow cytometry alone cannot
conclusively identify apoptotic cells; however, parallel morphological
studies using fluorescence microscopy confirmed the increase in the
number of cells with nuclear condensation and fragmentation, which is
characteristic of apoptotic morphological changes. This correlated well
with the increase in sub-G1 signal, which
suggested that the sub-G1 population did indeed
represent apoptotic cells. There is always a small percentage of
apoptotic cells in any culture. This is normal and is not related to
the presence of DMSO at the concentrations used. However, when cells
were treated with PhIP for 40 h and then cultured in PhIP-free
medium for a further 4 days, a population of
sub-G1 cells emerged that increased with
increasing PhIP concentration (Fig. 1, q-t,
and Table 1
).
The induction of apoptotic cells was coincident with S-phase arrest
(Fig. 1, q-s,
and Fig. 2
). To illustrate this point, Fig. 2
is constructed from the data
provided in Fig. 1
. At the highest concentrations of PhIP (5 and 10
µg/ml), the majority of cells were undergoing apoptosis (Fig. 1t,
Table 1
, and Fig. 2
). In contrast, cells in
sub-G1 comprised less that 5% in vehicle-treated
(DMSO) TK6 cells (Fig. 1, f, k, p, u).
It seems that treatment with PhIP for 20 h caused
reversible cell cycle damage, whereas a 40-h treatment caused a severe
cell cycle disruption that resulted in cells being arrested in S phase
and subsequently excluded by apoptotic cell death.
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shows that most cells were in G1 before
treatment. After 20 h, cells treated with DMSO progressed into S
phase and G2-M (Fig. 3b),
whereas after 20 h treatment with PhIP (10 µg/ml), more cells
accumulated in S-phase and less cells progressed into
G2-M, which indicated an S-phase delay. Control
cultures that were untreated showed a very similar cycle distribution
to that observed with DMSO treatment. At the 20-h time point, the
numbers of cells in G1 was similar in both DMSO-
and PhIP-treated samples (Fig. 4)
, which suggested that there was no delay in the cell cycle progression
through M to G1 and from G1
to S phase. At 40 h, more cells were distributed in S phase plus
G2-M in the sample treated with PhIP (10 µg/ml)
than those treated with DMSO ([Fig. 3
(compare c with
h) and Fig. 4
]. The cell-cycle delay effects that were
induced by a 40-h treatment with PhIP were still evident up to 4 days
after compound withdrawal [Fig. 3
(compare d, e,
f with i, j, k].
Interestingly, in this G1-rich population, only a
small proportion of the cells were in sub-G1,
even at the highest concentration of PhIP (Fig. 3k),
which
is in contrast with the asynchronized cell population (Fig. 1t).
From these observations, it seems that the early
response of cells to PhIP treatment differs with the phase of the cell
cycle. In G1 cells, the response is mainly
S-phase delay, whereas post-G1 cells undergo
S-phase arrest and subsequently apo-ptotic cell death.
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illustrates the flow cytometric bivariate distribution of TK6 cells in
which cellular DNA content was plotted versus cyclin A
content. Cyclin A protein started to accumulate in early- S-phase cells
and reached a maximum level in G2-M cells. This
expression pattern was observed in cells treated with both DMSO and
PhIP. However, when S-phase delay was caused by PhIP treatment, a
decrease in cyclin A was observed. Fig. 6
is representative of an experiment repeated several times. For clarity,
it is constructed from the data presented in Fig. 3
and illustrates
that PhIP treatment for 20 h caused a 17% increase in S-phase
cells (Fig. 3)
, whereas the mean fluorescence intensity of FITC,
indicative of the level of cyclin A expression, was decreased by 11%
compared with that after DMSO treatment. This result suggests that the
S-phase delay and the blockage of S phase to G2-M
transition induced by PhIP may be caused by decreased levels of cyclin
A.
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. The colony formation rate in cells treated with PhIP for 20 h
was not different from the control (Fig. 7)
. However, at this 14-day
time point, colonies formed in the plates treated with PhIP for 40 h were much smaller and not as readily distinguishable as those in
untreated plates, which indicated a delayed cell growth. Thus, for
these cells, the colony formation rate was reassessed on day 21, when
colony size was comparable with that of the day-14 untreated plates.
Untreated cells remained >95% viable at day 21. Fig. 7
shows that a
40-h treatment with PhIP decreased the colony formation rate in a
concentration-dependent manner. These results are consistent with the
flow cytometry study and indicate that the PhIP treatment may have
delayed the cell growth and/or lowered the cell viability.
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). However, after 40 h treatment with PhIP, the frequency of
hprt mutation significantly increased in a dose dependent
manner (Fig. 8)
. In cultures where the highest concentration of PhIP
was used (10 µg/ml), the incidence of mutation although significantly
higher than the DMSO control, was decreased (Fig. 8)
, presumably due to
PhIP induced cell death.
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DISCUSSION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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We also studied the early cellular responses induced by PhIP treatment in an attempt to define the events associated with genotoxicity. Significantly, we showed by flow cytometry that PhIP induced S-phase cell cycle delay, which started about 20 h after treatment and was clearly evident after 40 h, particularly at the highest concentration of PhIP (10 µg/ml). Propidium iodide-stained cells, viewed under fluorescence microscopy, showed a small percentage of apoptotic cells to be present at these time points. Intriguingly, cells that were treated with PhIP for 20 h resumed normal cell cycle when the compound was withdrawn, whereas the damage caused by 40-h PhIP treatment seemed to become more pronounced 4 days after PhIP withdrawal, and the concentration-dependent S-phase delay correlated well with the induction of apoptosis. At the highest concentration of PhIP, apoptosis was higher than the number of cells in S phase. It seems that the S-phase checkpoint was activated shortly after PhIP treatment; cells either progressed through the cell cycle or underwent apoptotic cell death. In accord with the effect on cell cycle, the time-and dose-dependent inhibition of cell growth by PhIP was also detected by a colony formation assay. These results indicate that two mechanisms are involved in cell growth inhibition induced by PhIP: (a) transient S-phase delay, which could not be assessed by colony formation assay; and (b) prolonged cell cycle arrest followed by apoptotic cell death, resulting in decreased colony formation.
Many DNA-damaging agents, including mutagens and carcinogens, induce the arrest of cell cycle at G1, S-phase, and G2 checkpoints, followed by apoptotic cell death (31 , 32) . Thus, apoptosis seems to be a useful marker for the measurement of genotoxicity. Cell cycle checkpoints may function to ensure cells have time for DNA repair (33 , 34) , whereas apoptotic cell death may function to eliminate irreparable or unrepaired damaged cells (35) . The fate of cells with DNA damage either to undergo apoptosis or to survive seems to be dependent on the intensity of DNA damage and the ability to repair DNA. Some DNA damage can be repaired completely, whereas severe damage may elicit apoptotic cell death. TK6 cells have a doubling time of about 16 h under the culture conditions we described; therefore, PhIP exposure for about one doubling time (20 h) may have resulted in low levels of DNA damage, which can be effectively repaired with little effect on cell cycle. However, if the exposure is extended to more than two doubling times (40 h), the DNA damage may accumulate to exceed a threshold for the activation of apoptosis.
Recent studies have highlighted the correlation between the chemical potential for the induction of apoptosis and carcinogenesis (36 , 37) . It has been reported that a single exposure of PhIP (100 mg/kg body weight) induced apoptosis within 24 h in rat colonic epithelium (38) , the site of PhIP-induced tumors. In concordance with this observation, our in vitro study demonstrated that PhIP induces apo-ptosis and also increases the rate of gene mutations in asynchronized TK6 cells.
Studies using G1-enriched cell populations demonstrated that at 20 h, samples treated with DMSO and PhIP contained similar numbers of G1 cells, which indicated that PhIP had not induced G1 blocking. Instead, at this time point, S-phase delay was evident. The difference in the number of G1 cells at 40-h treatment and on the following day (when the cells were incubated in fresh medium) could be due to the initial S-phase delay induced by PhIP. After PhIP withdrawal for 2 days, G1 cells in both samples reached the same number, and cell cycle profiles showed that they were at the same stage of the cell cycle. Interestingly, only a small proportion of cells in the G1-enriched population were eliminated by apoptosis over the course of the experiment. These observations suggest that PhIP could also delay G1 cells in S phase. However, they appear to be more resistant to apoptotic cell death. The phenomenon that G0-G1 phase cells are more resistant to apoptosis has been reported by others using different cell types and apoptotic agents (39 , 40) . The damaged cells could, therefore, progress into the next cycle, which would increase the potential for mutation.
TK6 cells express wild type of p53 (41) , a protein that mediates G1 checkpoint and apoptosis. The lack of G1 blocking in cells treated with PhIP indicates that the G1 checkpoint is not activated. A similar effect has also been demonstrated by Little et al. (42) using ionizing radiation. We found that PhIP treatment decreased the level of cyclin A, a protein that is associated with cdk2 or cdc2 and binds to the transcription factor E2F. E2F is responsible for the control of S-phase progression and G2-M transition (43, 44, 45) . The decrease in cyclin A level may result in a decrease in cdk2 and cdc2 activity, which leads to S-phase cell cycle arrest and subsequent downstream events. Evidence for such a mechanism exists because S-phase-related apoptosis was induced in NIH3T3 fibroblasts by disassociating E2F1 with cyclin A-cdk2 (24) , which proved the importance of cyclin A in the maintenance of S-phase checkpoint.
The finding that cells treated with PhIP apparently elude G1 and G2-M checkpoints but can be stalled at S phase is significant. Activation of the S-phase checkpoint mechanism may allow the cell to accumulate genetic damage, and DNA replication will continue using a damaged template; alternatively, it may permit the cell to activate DNA repair mechanisms. Under these conditions, the dynamic processes of DNA damage and repair are competing for the same template. If damage can be repaired, which seems to be the case after 20 h incubation with PhIP, then the cell can continue to division with minimal genetic damage. If, however, DNA damage predominates, as seems likely when PhIP is incubated with cells for 40 h, then cells will either be eliminated via apoptosis or proceed to replication carrying excessive levels of DNA damage, which would result in an increased chance of mutation. If similar mechanisms exist in vivo, then avoidance of G1 and G2-M checkpoints could increase the likelihood of neoplastic disease and tissues with a population of dividing cells might be expected to be more susceptible to PhIP-induced genetic damage. It is, therefore, perhaps no coincidence that, in rodent experiments, PhIP primarily induces tumors of the colon, mammary gland, and bone marrow tissues, all of which include a population of cells actively undergoing replication.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Supported by funds provided by the Ministry of
Agriculture Fisheries and Food, United Kingdom. 
2 To whom requests for reprints should be
addressed, at Molecular Toxicology, Imperial College School of
Medicine, Sir Alexander Fleming Building, South Kensington, London, SW7
2AZ United Kingdom. Phone: 0171-594-3188; Fax: 0171-594-3169; E-mail: n.gooderham{at}ic.ac.uk
3 The abbreviations used are: HCA,
heterocyclic amine; PhIP,
2-amino-1-methyl-6-phenylimidazo[4,5-b]pyridine;
hprt, hypoxanthine-guanine phosphoribosyl transferase;
6-TG, 6-thioguanine.
Received 6/11/99. Accepted 1/ 5/00.
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REFERENCES |
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) or 40 h (
) in the presence of irradiated
XEMh1A2-MZ cells. Cells were cultured for 6 days in PhIP-free medium
before they were moved to selective medium for colony formation assay.
The mutation frequency was the ratio of cloning efficiency of
nonselective culture:cloning efficiency of selective culture. Values
are mean ± SD of 3 independent experiments (4 plates
per experiment). Significantly different from control (DMSO);
***, P < 0.001; **,
P < 0.05.