| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
Experimental Therapeutics |
Department of Medicine and the Cancer Center, University of California, San Diego, La Jolla, California 92093-0058
 |
ABSTRACT |
|---|
 Top ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Among the types of defects that can cause genomic instability, the loss of MMR3 is of particular interest with respect to resistance to chemotherapeutic agents. MMR is a postreplicative DNA repair process that corrects single-base mismatches and small mismatched loops in the daughter strand of newly replicated DNA (6) . Failure to correct such mismatches results in an increased mutation rate, most readily apparent as instability in the length of microsatellite sequences. Loss of MMR because of mutation of MSH2 or MLH1 underlies the majority of cases of hereditary nonpolyposis colon cancer (8 , 9) and is also common in a variety of sporadic cancers including endometrial, ovarian, breast, prostate, lung, and pancreatic cancer (10) . In addition to increasing the rate of mutation throughout the genome, loss of MMR produces drug resistance directly by impairing the ability of the cell to detect adducts in its DNA that mimic base mismatches. Loss of MMR results in high-level resistance to the antimetabolite 6-thioguanine (11) , moderate levels of resistance to the methylating agents such as N-methyl-N'-nitro-N-nitrosoguanidine (7) and temozolomide (12) , and low-level resistance to DDP and carboplatin (13) in cell lines cultured in vitro. At concentrations attainable in patients, these agents cause cell death by apoptosis. Triggering apoptosis requires the cell to be able to recognize the presence of the damage in DNA, and the current hypothesis is that loss of MMR results in impaired adduct detection or lack of attempted repair and, thus, a diminished pro-apoptotic signal (7) .
p53 is another protein critical to the maintenance of genomic integrity, particularly after genotoxic stress. Increases in p53 after cellular injury mediate Gl checkpoint activation and enhanced DNA repair. Under circumstances where damage is extensive, p53 plays a direct role in triggering apoptosis (14) . Cells lacking p53 function are genetically unstable and are predisposed to gross genomic alterations such as gene amplification, aneuploidy, translocation, and deletions. p53 dysfunction is associated with increased tumorigenesis in p53 knockout mice (15) , and p53 is mutated in a very large fraction of human cancers (16) .
DDP is widely used for the treatment of a variety of solid tumors. The most abundant lesions produced in DNA are intrastrand cross-links, which are believed to account for both the cytotoxicity and mutagenicity of the drug. Its mutagenic potential has been well documented in both bacterial (17 , 18) and mammalian cells (19 , 20) . We have established that loss of MMR in the HCT116 cells causes low-level resistance to the cytotoxic effects of DDP (13) and also renders these cells hypersensitive to its mutagenic effects (21 , 22) . Because loss of p53 in human tumors is even more common than loss of MMR, it is of interest to determine how loss of these two genomic guardians interact. The aim of the current study was to determine whether loss of MMR function alters the effect of loss of p53 on parameters likely to be important to the development of drug resistance in the HCT116 model system.
|
MATERIALS AND METHODS |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Reagents.
Cisplatin and paclitaxel were gifts from Bristol-Myers Squibb
(Princeton, NJ). A stock solution of 1 mM cisplatin in
0.9% NaCl was stored in the dark at room temperature. Paclitaxel was
dissolved in DMSO, diluted with saline to form a stock solution of 5
µM, and stored at -20°C. Topotecan was purchased from
Smith Kline Beecham Pharmaceuticals (King of Prussia, PA), dissolved in
deionized water, and stored as a 10 µM stock solution at
-20°C. The clinical formulation of gemcitabine was purchased from
Eli Lilly and Co. (Indianapolis, IN) and was diluted directly in tissue
culture medium. The clinical formulation of etoposide was obtained from
Bristol-Myers Laboratories (Syracuse, NY) and was diluted directly in
tissue culture medium. 6-Thioguanine was purchased from Sigma Chemical
Co. (St. Louis, MO) and dissolved in 0.2 N sodium hydroxide
to form a 20 mM stock solution and stored at
-20°C.
Clonogenic Assay.
Clonogenic assays were performed by seeding 250 cells into 60-mm
plastic dishes in 5 ml of complete media. After 24 h, appropriate
amounts of the drugs were added to the dishes, and the cells were
exposed for 24 h (6-thioguanine, etoposide, and paclitaxel) or
1 h (all of the other drugs). Thereafter, the cells were washed,
and fresh drug-free medium was added. Colonies of at least 50 cells
were scored visually after 8–10 days. Each experiment was performed a
minimum of three times using triplicate cultures for each drug
concentration. IC50 values were determined using
log-linear interpolation.
Mutant Frequency Assay.
All of the HCT116 sublines were grown in
hypoxanthine-aminopterin-thymidine medium containing 0.4
µM aminopterin, 16 µM thymidine, and 100
µM hypoxanthine for a minimum of 14 days and then were
exposed for 1 h to increasing concentrations of DDP. Thereafter,
the cells were washed twice and recultured in regular medium for 8 days
during which the cultures were split 2:1 as needed to keep them from
becoming confluent. All of the cells were then trypsinized and seeded
into each of 10 100-mm tissue culture dishes at 100,000 cells/dish in
the presence of 20 µM 6-thioguanine. At the same time,
aliquots of 250 cells were seeded into each of three 60-mm dishes in
drug-free medium for determination of cloning efficiency. After 14
days, colonies were counted after staining with 0.1% crystal violet.
The procedure for determination of the frequency of mutation to the
other drugs used in this experiment was the same except that the cells
were not grown in hypoxanthine-aminopterin-thymidine medium before the
start of the experiments. A concentration of drug that resulted in a
cloning efficiency of approximately 0.0002% was added to identify the
number of resistant clones. These concentrations were 20
µM 6-thioguanine, 10 µM etoposide, 200
nM topotecan, 100 nM gemcitabine, and 20
nM paclitaxel. The frequency of resistant variants was
calculated as follows: variant frequency = a/(b x 106), where a is the number of
colonies present in the 10 drug-treated dishes and b is the
cloning efficiency. Each experiment was performed a minimum of
three times, and the data are presented as mean ± SD.
Shuttle Vector Mutation Assay.
The pZCA29 vector (27)
was obtained from Dr. T. M. Runger
(University of Gottingen, Gottingen, Germany). Four million
cells were transfected with 2 µg of pZCA29 by electroporation on day
1. Replicated pZCA29 was recovered from aliquots of the transfected
cells on days 3, 5, 7, 9, and 11 by a rapid alkaline lysis procedure.
For the DDP treatment experiments, the cells were treated with 25
µM DDP for 1 h 2 days after transfection, and the
vector was harvested on days 3, 5, 7, 9, and 11. Any pZCA29 that failed
to replicate in the mammalian cells is characterized by the bacterial
pattern of methylation. Such unreplicated plasmid DNA was removed by
digestion with DpnI, which cleaves the methylated DNA.
Escherichia coli XL1-Blue MRF’ (Stratagene) was transformed
with the recovered pZCA29 and then selected on LB agar plates
containing 5-bromo-4-chloro-3-indolyl-ß-galactosidase,
isopropyl-ß-D-thiogalactoside, and ampicillin.
Bacterial transformations were performed in triplicate for each of two
to three independent samples of pZCA29 recovered at each time point.
The mutant frequency was calculated as the mean of the total number of
blue colonies divided by the mean of the total number of colonies.
Flow Cytometry.
Subconfluent cultures growing in 10-cm tissue culture dishes were
exposed to 25 µM DDP for 1 h. At 1, 2, 3, 4, 5, 6,
and 7 days after DDP treatment, cells were harvested by trypsinization,
washed twice with ice-cold PBS, fixed in ice-cold 70% ethanol, treated
with RNase (Sigma) at 37°C for 30 min, and stained with 50 µg/ml
propidium iodide (Sigma). After a 30-min incubation on ice, the cells
were analyzed on a FACScan flow cytometer (Becton Dickinson, San Jose,
CA) using the FlowJo cell cycle analysis software (Tree Star, Inc., San
Carlos, CA) and the "Watson Pragmatic" model.
Western Blotting.
Cells were collected at times from 1 to 7 days after a 1-h treatment
with 25 µM DDP and lysed in 100 µl of lysis buffer [10
mM Tris-HCl (pH7.4), 150 mM NaCl, 5
mM EDTA, 1% Triton X-100, 5 mM dithiothreitol,
1 mM sodium vanadate, 0.1 mM
phenylmethylsulfonyl fluoride, and 5 mM aminocaproic acid]
for 30 min on ice. The insoluble material was removed by centrifugation
at 14,000 x g for 20 min at 4°C. Protein
(10 µg) from each sample was electrophoresed through 10–20%
SDS-PAGE and transferred to a polyvinylidene difluoride membrane
(Immobilon P; Millipore, Bedford, MA). The membranes were blotted with
monoclonal antibodies specific for p53 (Santa Cruz Biotechnology, Santa
Cruz, CA). After application of a horseradish peroxidase-coupled
secondary antibody, reactive proteins were visualized with enhanced
chemiluminescence (Amersham, Arlington Heights, IL).
Platinum-DNA Adduct Formation and Repair.
The binding of platinum to DNA was determined in whole cells exposed to
DDP. For dose-response studies, cells at 80% confluence in T-150
flasks were incubated in fresh medium containing 0–200
µM drug for 1 h. The cells were then trypsinized,
washed three times with PBS, and incubated overnight at room
temperature in a lysis buffer containing 0.67% Triton X-100, 2.6
M NaCl, 133 mM EDTA, and 2.6
M Tris-HCl (pH 8.0). DNA was isolated by phenol-chloroform
extraction and dissolved in TE buffer (pH 8.0). Aliquots of the
DNA were digested in 1 M HCl at 75°C for 2 h, and
the hydrolysate was used for the quantitation of platinum by flameless
atomic absorption spectrophotometry (Perkin-Elmer Model 2380). These
experiments verified that platinum-DNA adduct levels were a linear
function of DDP concentration in all of the four HCT116 sublines.
For measurement of the disappearance of platinum from DNA, subconfluent cells in T-150 flasks were treated for 1 h with DDP concentrations that yielded the same initial adduct level, and the pg platinum/µg DNA was measured at 0, 3, 6, 12, and 24 h after the drug treatment.
Statistics.
All of the data were analyzed by use of a two-sided paired Student’s
t test with the assumption of unequal variance.
|
RESULTS |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
, and the ratios of IC50 values are summarized in
Table 2
. Loss of p53 function in MMR-proficient HCT116+ch3 cells conferred
statistically significant resistance only for topotecan. However,
disruption of p53 in MMR-deficient HCT116 cells produced significant
degrees of resistance to gemcitabine, 6-thioguanine, and etoposide by a
factor of 2.7-, 3.3- and 4.4-fold, respectively, but paradoxically
rendered the cells hypersensitive to topotecan and DDP by factors of
2.5- and 3.3-fold, respectively. Thus, the effect of disabling p53, MMR
function, or both together was unique to each drug, consistent with the
fact that each has a different mechanism of action. This suggests that
the interaction between p53 and MMR with respect to cytotoxicity varies
depending on the type of injury inflicted.
|
|
, the parental cells and the heterozygote with one deleted
p53 allele demonstrated similar sensitivity with
IC50 values of 24.4 µM
and 18.4 µM, respectively. However, the
homozygote in which both p53 alleles were disrupted was
markedly more sensitive with an IC50 of 5.7
µM, thus confirming the results obtained with
the E6-expressing cells. This finding is in good agreement with prior
observations showing that knockout of p53 makes these human colorectal
cancer cells more sensitive to apoptosis induced by other agents
capable of inducing DNA damage such as doxorubicin (26)
and 
-radiation (31)
.
|
presents these slopes for 6-thioguanine, etoposide, topotecan,
gemcitabine, and paclitaxel for the four HCT116 sublines. On the basis
of the ratio of the slopes, as shown in Table 4
, loss of p53 in the MMR-proficient HCT116+ch3 cells had only a minor
effect on the ability of DDP to generate variants resistant to
6-thioguanine, etoposide, topotecan, gemcitabine, or paclitaxel. In
contrast, loss of p53 in the MMR-deficient HCT116 cells resulted in
larger increases in slope for all of these drugs. Stated the other way
around, loss of MMR in the p53-proficient HCT116+ch3 cells had a modest
impact, producing increases of 1.1–2.4-fold, whereas loss of MMR in
the p53-deficient HCT116+ch3/E6 cells had more profound effect and
increased the generation of resistant variants by 2.3–6.5-fold. These
results indicate that p53 and MMR both operate to prevent the emergence
of resistant clones after exposure to DDP and that they do so by at
least partially independent mechanisms because loss of both functions
produced a greater effect than loss of either alone.
|
|
shows that when both p53 alleles were deleted from the
parental cells, DDP was 3.9-fold (P < 0.01)
more effective at generating 6-thioguanine-resistant variants.
Interestingly, when one p53 allele was deleted there was
little change in sensitivity at low DDP concentrations; however, at
higher DDP concentrations there was nearly the same increase in
resistant variants generated as was observed when both alleles were
deleted. This gene dosage effect is consistent with the hypothesis that
at low adduct burdens the amount of p53 generated by a single allele is
sufficient to guard against mutation at most sites, but at high adduct
burdens it is inadequate.
|
Fig. 3A
shows the spontaneous mutant frequencies observed after
replication of the pZCA29 vector in the HCT116+ch3
(MMR+/p53+), HCT116+ch3/E6
(MMR+/p53-), HCT116
(MMR-/p53+), and HCT116/E6
(MMR-/p53-) cell lines
for various periods of time. These frequencies reflect the ability of
the cell to faithfully replicate the out-of-frame vector under basal
conditions in the absence of any exogenous insult. For each cell line,
the number of mutations increased as a function of the time during
which the vector was allowed to replicate in the tumor cell.
Differences between the cell lines were already apparent after just 3
days of vector replication. The mutant frequency was lowest for the MMR
and p53-proficient HCT116+ch3 cell line. Loss of either p53 function or
MMR function alone resulted in a small increase in the number of
mutants; the increases were 1.2- and 1.7-fold, respectively
(P > 0.05 for all of the time points
relative to the MMR+/p53+
cells). However, the greatest increase was observed in the cells that
had lost both p53 and MMR function (5.1-fold). This increase was
statistically significant (P < 0.05) for
comparison with all of the other three sublines at all of the time
points measured. Thus, the pZCA29 vector detected the type of genomic
instability produced by loss of either p53 or MMR, and this type of
instability was augmented in cells burdened by loss of both functions.
|
shows that treatment of cells with 25
µM DDP for 1 h starting 48 h after
introduction of the vector into the tumor cells increased the number of
pZCA29 mutants in all of the four cell lines. On the basis of the
slopes of the curves, loss of p53 permitted DDP exposure to generate an
average of 1.2-fold more mutants (P = 0.67),
whereas loss of MMR function permitted it to generate an average of
1.7-fold more mutants (P = 0.13). Loss of
both p53 and MMR function in the HCT116/E6 cells resulted in 5.1-fold
more mutants (P = 0.03). Loss of p53 function
had a larger effect in MMR-deficient cells (2.9-fold;
P = 0.05) than in MMR-proficient cells
(1.2-fold; P = 0.67). Likewise, loss of MMR
had a larger effect in p53-deficient cells (4.3-fold;
P = 0.04) than in p53-proficient cells
(1.7-fold; P = 0.13). Thus, DDP was much more
effective at producing substitution and/or deletion mutations when both
p53 and MMR function were lost.
To confirm the results obtained when expression of E6 was used
to disable p53 function, the pZCA29 vector was used to measure the
basal and DDP-induced frequency of mutations in the HCT116 sublines in
which one or both p53 alleles had been deleted. The basal
mutant frequencies after passage of the pZCA29 vector through the
HCT116 p53+/+, p53+/-, and
p53-/- cell lines are shown in Fig. 4A
. The p53+/- cells replicated the
vector as faithfully as wild-type cells over the period tested.
However, loss of both p53 alleles rendered the vector
replication error-prone by a factor of 1.9-fold (P = 0.12). Fig. 4B
shows that when cells were treated
with 25 µM DDP for 1 h starting 48 h
after vector introduction, the number of mutants produced was increased
in all of the three cell lines. On the basis of the ratio of the
slopes, DDP generated 1.5-fold (P = 0.19) and
2.9-fold (P = 0.04) more mutants,
respectively, in the p53+/- and
p53-/- cells compared with the parental
p53+/+ HCT116 cells. Thus, the simple repetitive
sequences in pZCA29 were hypermutable after DDP exposure in the cells
in which p53 function was inactivated.
|
, in the p53- and MMR-proficient HCT116+ch3 cells, DDP induced a marked
G2-M phase arrest that peaked at 24 h at
which point 65.1% of the DDP-treated cells were in
G2-M compared with 29.5% of the untreated
control population. The G2-M arrest gradually
resolved over the ensuing days such that the cell cycle phase
distribution had returned to normal by 7 days after DDP exposure. Loss
of p53 alone resulted in a greater depletion of cells from
G1 at early time points such that at 24 h
after DDP exposure only 6.6% of the cells were in
G1 compared with 23.4% of cells with intact p53
function. The peak accumulation of cells in the
G2-M phase was also increased (to 63.8%) and
delayed to 48 h compared with the p53-proficient cells. Loss of
MMR-function alone also resulted in a greater loss of cells from
G1 over the first 24 h. However, the largest
effect was a more rapid resolution of the G2-M
phase accumulation consistent with failure of the cells to remain
arrested in G2-M. When both MMR and p53 functions
were disabled, the perturbations produced by loss of p53 predominated
over those produced by loss of MMR. There was excessive depletion of
cells from G1 at all of the time points examined
and a substantial delay in the resolution of the
G2-M phase arrest. Thus, loss of either p53 or
MMR function alone produced unique patterns of checkpoint failure, and
the greatest difference from the pattern observed in the fully
proficient cells occurred when both functions were lost.
|
, in the p53+/MMR+ cells,
p53 levels were increased by 1 day after exposure, and high levels
persisted for up to 6 days. This response was blunted in cells in which
MMR had been disabled, with both the magnitude and duration of the
change in p53 levels being reduced. As expected based on the ability of
E6 to mediate the degradation of p53, although p53 was detectable in
the E6-expressing HCT116+ch3/E6 and HCT116/E6 cells, the levels did not
change significantly after DDP exposure, and they were consistently
below those found in the non-E6-expressing cells (data not shown).
Thus, loss of MMR had a substantial effect on the extent to which
DDP-DNA adducts generated the signals responsible for the stabilization
of p53.
|
shows that there were substantial differences in the kinetics of
platinum disappearance from DNA among the four sublines. By 24 h
after the start of the 1-h drug exposure, only 5% of the platinum
remained in the DNA of the HCT116+ch3 cells. Loss of either p53 or MMR
function alone reduced the disappearance rate such that by 24 h
20% and 17%, respectively, of the platinum remained in the DNA. Loss
of both p53 and MMR resulted in further slowing of platinum
disappearance with persistence of 39% of the initial load in the DNA
at 24 h. To better quantify the platinum clearance from DNA among
the four sublines, the area under the curve for the percentage of
platinum-DNA adducts remaining over time (percentage x h) was calculated using trapezoidal rule. The area under the curve for
HCT116+ch3, HCT116+ch3/E6, HCT116, and HCT116/E6 cells was 619.5,
1042.5, 1141.5, and 1537.5 (percentage of platinum remaining x h), respectively. This indicates that loss of p53 alone
resulted in a 1.7-fold increase in total adduct exposure, whereas loss
of MMR alone produced a 1.8-fold increase, and loss of both caused a
2.5-fold increase.
|
As shown in Table 5
, total platinum/µg DNA at the end of a 1-h exposure increased
linearly with DDP concentration for all of the four sublines, and the
adduct potency was remarkably constant at different DDP concentrations
within each subline. Loss of p53 function alone increased adduct
tolerance by 2.0-fold (P = 0.0001), whereas
loss of MMR alone increased tolerance by a factor of 2.7-fold
(P = 0.0083). Thus, both functional deficits
permitted cells to survive with a higher adduct load in their DNA.
Interestingly, instead of generating an even higher level of tolerance,
loss of both MMR and p53 function resulted in only a 1.8-fold increase
in tolerance (P = 0.0144). This suggests
that, with respect to adduct tolerance, both p53 and MMR may be
operating through the same mechanism.
|
|
DISCUSSION |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Loss of MMR results in genomic instability characterized by small insertion and deletion mutations in repetitive sequences throughout the genome. Our prior work (21 , 22) demonstrated that loss of MMR rendered cells hypersensitive to the ability of DDP to generate clones resistant not only to 6-thioguanine but also to etoposide, topotecan, gemcitabine, and paclitaxel. Loss of MMR is particularly important with respect to the emergence of drug resistance because MMR-deficient cells are not only hypersensitive to exogenous and endogenous mutagens (35 , 36) but also resistant to DDP because of an apparent reduced ability of the cell to sense the presence of adducts in DNA (37 , 38) . It has been well documented, using both in vitro and in vivo models, that the degree of resistance is great enough so that subsequent treatment with DDP enriches for these genomically unstable cells (39) .
The effect of the loss of p53 and MMR function alone on chemotherapeutic agent cytotoxicity appears to be unique to the cell line under study, and both increased and decreased sensitivities have been observed in different model systems (13 , 26 , 40, 41, 42, 43) . The results reported here are in general agreement with prior studies in colon cancer cell lines (13 , 26) . In the colon carcinoma cells used in this study, the effect of loss of both p53 and MMR varied among the drugs tested. In the case of 6-thioguanine, etoposide, and gemcitabine, loss of both p53 and MMR incrementally increased the level of resistance over that observed with loss of either function alone. Thus, it would be reasonable to expect that treatment with any of these agents would enrich the tumor for doubly deficient cells.
Specifically with respect to changes in sensitivity to the cytotoxic effect of DDP, some investigators have reported that loss of p53 function renders cells hypersensitive whereas others have found that it renders them resistant (summarized in Ref. 44 ) even in cells that are likely to have intact MMR (42 , 45) . In the model system used in the current study, loss of p53 had relatively little effect in MMR-proficient cells but conferred substantial hypersensitivity in MMR-deficient cells, a finding that is consistent with prior observations by Vikhanskaya et al. (43) . Although a central role for p53 in mediating activation of the caspase cascade after DNA damage has been extensively reported, and DDP evokes a large increase in p53 level in the p53+/MMR+ subline, any proapoptotic effect of this change is apparently offset in these cells by other antiapoptotic roles played by this protein. In the absence of functional MMR, these cells appear to be more dependent on this protective effect of p53, as the impact of the loss of p53 was substantially larger in MMR-deficient than MMR-proficient cells. This finding is consistent with experiments in other cellular systems. Brown et al. (28) reported that transfection of a dominant negative mutant p53 in A2780 cells (MMR proficient) did not significantly change DDP-induced cytotoxicity, whereas the same vector transfected into an MMR-deficient subline of A2780 induced a significant increase in DDP sensitivity. In addition, by comparing E6-transfected clones obtained from HCT-116 or MCF cells, it was noted that disruption of p53 in MMR-deficient cells (HCT116) greatly enhanced sensitivity to DDP (43 , 46) , whereas in MCF-7 cells in which MMR function was normal, the effects were much diminished (41) . Therefore, our findings extend observations made in other cell lines and further support the hypothesis that p53 cooperates with MMR in determining the cellular sensitivity to DDP.
A clear interaction between loss of p53 and loss of MMR was apparent also with respect to the mutagenic potential of DDP. Loss of both p53 and MMR function rendered the cells substantially more sensitive to the ability of DDP to generate variants resistant to a variety of other drugs than loss of either p53 or MMR alone. Although we did not document that the resistant clones were true mutants, the presence of such clones capable of withstanding very high level exposures to these drugs is likely to contribute to the emergence of drug resistance. This finding brings into sharp focus the concern that initial treatment with agents such as DDP, although it may help reduce tumor burden, also sows the seeds that will eventually cause treatment failure by generating clones within the tumor that are resistant to many other types of drugs. The specific mechanisms by which DDP generates these resistant variants are not yet understood. However, it has been demonstrated that single mutations in topoisomerase II can produce resistance to etoposide (47) . In the case of topotecan and gemcitabine, it has already been established that resistance can result from mutations in the topoisomerase I (48) and deoxycytidine kinase (49) gene, respectively. As for paclitaxel, the target of which is ß tubulin, the role of single mutations in mediating resistance has not been as clearly established. However, previous studies have documented that paclitaxel-resistant variants arise at a rate as high as that for etoposide (50) . That p53 and MMR both function to modulate the ability of DDP to produce mutations was documented further by the experiments with the pZCA29 vector. An increased frequency of small insertion/deletion mutations is a feature of the genomic instability produced by the loss of either p53 or MMR (6 , 27 , 51) , and the pZCA29 vector proved capable of detecting these changes in the basal rate of mutation. It also effectively detected the mutagenic effect of DDP exposure in the p53+/MMR+ cells. Although loss of either p53 or MMR alone increased DDP-induced mutations, the observation that the loss of both functions produced a further increase argues that p53 and MMR operate in different pathways to offset the mutational risk posed by DDP adducts in DNA.
Previous studies (35) from this laboratory have demonstrated that p53 and MMR-deficient HCT116/E6 cells are resistant to exogenous H2O2 but hypersensitive to the ability of H2O2 to generate mutants resistant to 6-thioguanine and ouabain. The current study did not investigate the biochemical nature of the interaction between loss of p53 and MMR; i.e., it cannot be determined from the studies reported here whether the interaction is truly synergistic, only additive, or even partially antagonistic. However, it is clear that DDP is more cytotoxic and mutagenic when both functions are disabled. Caution is needed in interpreting the resulting effects of E6 expression and chromosome 3 transfer as being entirely because of the loss of p53 and complementation of MMR, respectively, because E6 can affect other cellular proteins, and transfer of the chromosome 3 also introduces other genes into the cell.
What is the mechanism by which loss of either p53 or MMR causes DDP to be more mutagenic? The results presented here suggest multiple effects. DDP exposure induced an increase in p53 and activated the Gl checkpoint in the p53+/MMR+ cells, and the differences observed in cell-cycle phase distribution when p53 function was lost are consistent with the premature release of cells into S phase. Mammalian DNA polymerases that can bypass adducts in DNA, but create mutations as they do so, have recently been identified (52 , 53) . Both p53 and MMR may play a role in preventing such mutagenic bypass replication (54 , 55) . Alternatively, if polymerase fidelity is influenced by MMR or p53, mutations may be introduced during the gap-filling step after processing of the adduct by one or another of the DNA repair mechanisms.
Among the known DNA repair mechanisms, nucleotide excision repair appears to be quantitatively the most important for removal of DDP adducts. Evidence for a direct role for p53 protein in this and other types of DNA repair is accumulating rapidly. Wang et al. (56) reported that p53 can bind to several proteins known to play central roles in nucleotide excision repair, including XPD (Rad3), XPB, and CSB, and Therrien et al. (57) reported direct involvement of p53 in both global and transcription-coupled nucleotide excision repair. Very recently, Tanaka et al. (58) reported that p53 regulates the transcription of a catalytic subunit of ribonucleotide reductase, an enzyme essential to the supply of deoxynucleotides for nucleotide excision repair. MMR also modulates at least one subtype of nucleotide excision repair, transcription-coupled repair. The observation that the kinetics of platinum disappearance from DNA are delayed is consistent with a cooperative interaction between p53 and MMR with respect to nucleotide excision repair. Although variable degrees of continued proliferation can contribute to differences in the disappearance of platinum from DNA, this is unlikely because of the high concentrations of DDP (100–200 µM) used in these kinetic studies.
One might have expected impairment of nucleotide excision repair because of loss of p53 or MMR function to result in hypersensitivity to the cytotoxic effect of DDP. Other genetic lesions that disable nucleotide excision repair clearly produce such an effect (59) . Instead, loss of p53 or MMR diminished the cytotoxic potency of DDP adducts by a factor of 2.0- and 2.7-fold, respectively. Thus, the reduced disappearance of adducts from DNA was offset in the p53- and MMR-deficient cells by improved adduct tolerance. This is of particular concern with respect to the emergence of drug resistance because the persistence of such adducts in DNA is likely to contribute additional mutations if the cell can complete another round of DNA synthesis. Such adduct tolerance may be important for other DNA-interacting drugs, although changes in sensitivity to drugs that do not produce adducts must be explained by effects on signaling or apoptotic mechanisms.
Mutations that disable p53 are found very frequently in human cancers (16) , often in association with tumor progression or high-grade malignancy (15) . Loss of MMR function is a less common but well-described phenomenon, particularly in colon and endometrial cancer (8 , 9) . Thus, there is a reasonable likelihood that many tumors contain at least a few cells in which both functions have been disabled. The results presented in this study suggest that these cells are particularly dangerous as potential mediators of the continued accumulation of somatic mutations that provide the heterogeneity that favors emergence of drug resistance.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
1 Supported in part by Grant CA78648 from the NIH.
This work was conducted in part by the Clayton Foundation for
Research-California Division. Drs. X. Lin, R. D. Christen, and S. B.
Howell are Clayton Foundation investigators. 
2 To whom requests for reprints should be
addressed, at Department of Medicine 0058, University of California,
San Diego, La Jolla, CA 92093. Phone: (619) 822-1110; Fax:
(619) 822-1111; E-mail: showell{at}ucsd.edu
3 The abbreviations used are: MMR, DNA mismatch
repair; DDP, cisplatin.
Received 6/ 9/00. Accepted 12/ 8/00.
|
REFERENCES |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
gene in patients with small lung cancer treated with etoposide.. Cancer Res., 56: 1232-1236, 1996.
This article has been cited by other articles:
|
|
 |
|
  L. P. Martin, T. C. Hamilton, and R. J. Schilder Platinum Resistance: The Role of DNA Repair Pathways Clin. Cancer Res., March 1, 2008; 14(5): 1291 - 1295. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. C. Young, A. M. Keuling, R. Lai, P. N. Nation, V. A. Tron, and S. E. Andrew The associated contributions of p53 and the DNA mismatch repair protein Msh6 to spontaneous tumorigenesis Carcinogenesis, October 1, 2007; 28(10): 2131 - 2138. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Zou, C. Tornos, X. Qiu, M. Lia, and R. Perez-Soler p53 Aerosol Formulation with Low Toxicity and High Efficiency for Early Lung Cancer Treatment Clin. Cancer Res., August 15, 2007; 13(16): 4900 - 4908. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. S. Hah, J. M. Mundt, H. M. Kim, R. A. Sumbad, K. W. Turteltaub, and P. T. Henderson From the Cover: Measurement of 7,8-dihydro-8-oxo-2'-deoxyguanosine metabolism in MCF-7 cells at low concentrations using accelerator mass spectrometry PNAS, July 3, 2007; 104(27): 11203 - 11208. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. J. Krieg, E. M. Hammond, and A. J. Giaccia Functional Analysis of p53 Binding under Differential Stresses. Mol. Cell. Biol., October 1, 2006; 26(19): 7030 - 7045. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Yu, M. A. Mallon, W. Zhang, R. R. Freimuth, S. Marsh, M. A. Watson, P. J. Goodfellow, and H. L. McLeod DNA Repair Pathway Profiling and Microsatellite Instability in Colorectal Cancer Clin. Cancer Res., September 1, 2006; 12(17): 5104 - 5111. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 X. Lin and S. B. Howell DNA mismatch repair and p53 function are major determinants of the rate of development of cisplatin resistance Mol. Cancer Ther., May 1, 2006; 5(5): 1239 - 1247. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 X. Lin, T. Okuda, J. Trang, and S. B. Howell Human REV1 Modulates the Cytotoxicity and Mutagenicity of Cisplatin in Human Ovarian Carcinoma Cells Mol. Pharmacol., May 1, 2006; 69(5): 1748 - 1754. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. W. Cheung, A. C.S. Chun, Q. Wang, W. Deng, L. Hu, X.-Y. Guan, J. M. Nicholls, M.-T. Ling, Y. Chuan Wong, S. Wah Tsao, et al. Inactivation of Human MAD2B in Nasopharyngeal Carcinoma Cells Leads to Chemosensitization to DNA-Damaging Agents. Cancer Res., April 15, 2006; 66(8): 4357 - 4367. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
X. Lin, J. Trang, T. Okuda, and S. B. Howell DNA Polymerase {zeta} Accounts for the Reduced Cytotoxicity and Enhanced Mutagenicity of Cisplatin in Human Colon Carcinoma Cells That Have Lost DNA Mismatch Repair Clin. Cancer Res., January 15, 2006; 12(2): 563 - 568. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S Kruger, A Bier, C Engel, E Mangold, C Pagenstecher, M von Knebel Doeberitz, E Holinski-Feder, G Moeslein, K Schulmann, J Plaschke, et al. The p53 codon 72 variation is associated with the age of onset of hereditary non-polyposis colorectal cancer (HNPCC) J. Med. Genet., October 1, 2005; 42(10): 769 - 773. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Okuda, X. Lin, J. Trang, and S. B. Howell Suppression of hREV1 Expression Reduces the Rate at Which Human Ovarian Carcinoma Cells Acquire Resistance to Cisplatin Mol. Pharmacol., June 1, 2005; 67(6): 1852 - 1860. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Yu, W. D. Shannon, M. A. Watson, and H. L. McLeod Gene Expression Profiling of the Irinotecan Pathway in Colorectal Cancer Clin. Cancer Res., March 1, 2005; 11(5): 2053 - 2062. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Fukushima, Y. Katayama, T. Watanabe, A. Yoshino, A. Ogino, T. Ohta, and C. Komine Promoter Hypermethylation of Mismatch Repair Gene hMLH1 Predicts the Clinical Response of Malignant Astrocytomas to Nitrosourea Clin. Cancer Res., February 15, 2005; 11(4): 1539 - 1544. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
F. Wu, X. Lin, T. Okuda, and S. B. Howell DNA Polymerase {zeta} Regulates Cisplatin Cytotoxicity, Mutagenicity, and The Rate of Development of Cisplatin Resistance Cancer Res., November 1, 2004; 64(21): 8029 - 8035. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. Bassett, N. M. King, M. F. Bryant, S. Hector, L. Pendyala, S. G. Chaney, and M. Cordeiro-Stone The Role of DNA Polymerase {eta} in Translesion Synthesis Past Platinum-DNA Adducts in Human Fibroblasts Cancer Res., September 15, 2004; 64(18): 6469 - 6475. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 F. Chen, O. K. Arseven, and V. L. Cryns Proteolysis of the Mismatch Repair Protein MLH1 by Caspase-3 Promotes DNA Damage-induced Apoptosis J. Biol. Chem., June 25, 2004; 279(26): 27542 - 27548. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Nakayama, A. Kanzaki, K. Terada, M. Mutoh, K. Ogawa, T. Sugiyama, S. Takenoshita, K. Itoh, N. Yaegashi, K. Miyazaki, et al. Prognostic Value of the Cu-Transporting ATPase in Ovarian Carcinoma Patients Receiving Cisplatin-Based Chemotherapy Clin. Cancer Res., April 15, 2004; 10(8): 2804 - 2811. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. L. Hayward, J. S. Macpherson, J. Cummings, B. P. Monia, J. F. Smyth, and D. I. Jodrell Enhanced oxaliplatin-induced apoptosis following antisense Bcl-xl down-regulation is p53 and Bax dependent: Genetic evidence for specificity of the antisense effect Mol. Cancer Ther., February 1, 2004; 3(2): 169 - 178. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
G. Samimi, N. M. Varki, S. Wilczynski, R. Safaei, D. S. Alberts, and S. B. Howell Increase in Expression of the Copper Transporter ATP7A during Platinum Drug-Based Treatment Is Associated with Poor Survival in Ovarian Cancer Patients Clin. Cancer Res., December 1, 2003; 9(16): 5853 - 5859. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. W. Robinson, M. M. Im, M. Ljungman, F. Praz, and D. S. Shewach Enhanced Radiosensitization with Gemcitabine in Mismatch Repair-Deficient HCT116 Cells Cancer Res., October 15, 2003; 63(20): 6935 - 6941. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A Brieger, J Trojan, J Raedle, G Plotz, and S Zeuzem Transient mismatch repair gene transfection for functional analysis of genetic hMLH1 and hMSH2 variants Gut, November 1, 2002; 51(5): 677 - 684. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. Raymond, S. Faivre, S. Chaney, J. Woynarowski, and E. Cvitkovic Cellular and Molecular Pharmacology of Oxaliplatin Mol. Cancer Ther., January 1, 2002; 1(3): 227 - 235. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Kondo, R. Safaei, M. Mishima, H. Niedner, X. Lin, and S. B. Howell Hypoxia-induced Enrichment and Mutagenesis of Cells That Have Lost DNA Mismatch Repair Cancer Res., October 1, 2001; 61(20): 7603 - 7607. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Cancer Research | Clinical Cancer Research |
| Cancer Epidemiology Biomarkers & Prevention | Molecular Cancer Therapeutics |
| Molecular Cancer Research | Cancer Prevention Research |
| Cancer Prevention Journals Portal | Cancer Reviews Online |
| Annual Meeting Education Book | Meeting Abstracts Online |
), p53+/- (•), and
p53-/- (
) HCT116 cells. Each point represents the mean
of three experiments with triplicate cultures. Bars,
SD.
), HCT116+ch3/E6 (•), and HCT116/E6 (
