a role for p53 in the frequency and mechanism of mutation, 2002
TRANSCRIPT
![Page 1: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/1.jpg)
Mutation Research 511 (2002) 45–62
Review
A role for p53 in the frequency and mechanism of mutation
Suzanne M. Morris∗
Division of Genetic and Reproductive Toxicology, National Center for Toxicological Research, 3900 NCTR Road, Jefferson, AR 72079, USA
Received 6 August 2001; received in revised form 3 December 2001; accepted 3 December 2001
Abstract
The tumor suppressor protein, p53, is often referred to as the guardian of the genome. When p53 function is impaired, its
ability to preserve genomic integrity is compromised. This may result in an increase in mutation on both a molecular and
chromosomal level and contribute to the progression to a malignant phenotype. In order to study the effect of p53 function
on the acquisition of mutation, in vitro and in vivo models have been developed in which both the frequency and mechanism
of mutation can be analyzed. In human lymphoblastoid cells in which p53 function was impaired, both the spontaneous and
induced mutant frequency increased at the autosomal thymidine kinase (TK) locus. The mutant frequency increased to a
greater extent in cell lines in which p53 harbored a point mutation than in those lines in which a “null” mutation had been
introduced by molecular targeting or by viral degradation indicating a possible “gain-of-function” associated with the mutant
protein. Further, molecular analysis revealed that the loss of p53 function was associated with a greater tendency towards
loss-of-heterozygosity (LOH) within the TK gene that was due to non-homologous recombination than that found in wild-type
cells. Most data obtained from the in vivo models uses the LacI reporter gene that does not efficiently detect mutation that
results in LOH. However, studies that have examined the effect of p53 status on mutation in the adenine phosphoribosyl
transferase (APRT) gene in transgenic mice also suggest that loss of p53 function results in an increase in mutation resulting
from non-homologous recombination. The results of these studies provide clear and convincing evidence that p53 plays a
role in modulating the mutant frequency and the mechanism of mutation. In addition, the types of mutation that occur within
the p53 gene are also of importance in determining the mutant frequency and the pathways leading to mutation. Published by
Elsevier Science B.V.
Keywords: p53; Mutant p53; Loss-of-function; p53 knockout
Abbreviations: IR, ionizing radiation; topo-I, topoisomerase-I;
topo-II, topoisomerase-II; HPV16E6, human papillomavirus Type
16 open reading frame E6; DDB2, DNA damage binding pro-
tein 2; ARF, alternate reading frame; TK, thymidine kinase; LOH,
loss-of-heterozygosity; HPRT, hypoxanthine-guanine phosphoribo-
syl transferase; 4-NQO, 4-nitroquinoline oxide; B[a]P, benzo(a)-
pyrene; DAP, 2,6-diaminopurine; APRT, adenine phosphoribosyl
transferase; pun, pink-eyed unstable mutation; NER, nucleotide
excision repair; 2-AAF, 2-acetylaminofluorene∗ Tel.: +1-870-543-7580; fax: +1-870-543-7136.
E-mail address: [email protected] (S.M. Morris).
1. Introduction
It is becoming increasingly accepted that the pro-
gression of mammalian cells towards malignancy is
an evolutionary process that involves an accumulation
of mutations on both the molecular and chromosomal
level. Inherent in models for malignant progression
is the concept that an initial mutation in an important
regulatory gene (protein) may be pivotal in this pro-
cess. Once the initial mutation has been introduced,
loss of normal gene function or the acquisition of
1383-5742/02/$ – see front matter. Published by Elsevier Science B.V.
PII: S1383 -5742 (01 )00075 -8
![Page 2: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/2.jpg)
46 S.M. Morris /Mutation Research 511 (2002) 45–62
deleterious functions may lead to additional mutations
furthering the malignant transformation of the cell.
A candidate for involvement in this process is the
tumor suppressor, p53. The p53 protein provides one
of the key regulatory elements monitoring genomic
integrity in mammalian cells and is involved in a
multiplicity of cellular functions. The p53 gene is
also recognized as the most commonly mutated gene
in human malignancies. Taken together, these factors
suggest that the role of “guardian of the genome” as-
cribed to p53 [1] may reflect its ability to reduce the
accumulation of mutation in the genome and inhibit
progression towards malignancy. An approach to test-
ing this hypothesis is to examine the role of mutations
in the p53 gene on the frequency and mechanism of
mutations in other genes.
2. Structure of p53
Knowledge of the structure of the p53 gene
provides some insight into the mechanism by which
mutations in p53 affect pathways important in the
mutational process. The human gene contains 11 ex-
ons that code for a protein 393 amino acids in length.
The p53 protein contains several functional domains
including the amino terminal transactivation domain
(amino acids 20–42), a proline-rich sequence con-
taining multiple copies of the PXXP sequence (amino
acids 60–97), a central DNA binding domain (amino
acids 100–293), a flexible linker region (amino acids
300–325), an oligomerization domain (amino acids
319–360) and a highly basic region of the C-terminus
(amino acids 363–393, reviewed in [2,3]). Sequences
found within the central DNA binding domain (exons
5–8) have been evolutionarily conserved across
species (reviewed in [2]).
The p53 protein undergoes extensive post-trans-
lational modification during an “activation” process.
Specific phosphorylation sites have been identified
and are located primarily in the amino and carboxy
terminal regions of the protein (reviewed in [3–12]).
The N-terminal domain functions primarily in the
transcriptional control activity of p53 whereas the C-
terminus affects the specific DNA binding of the
protein [4]. The amino terminal domain undergoes
phosphorylation by kinases that include casein kinase,
checkpoint kinase 1 and checkpoint kinase 2, DNA-
dependent protein kinase, ataxia telangiectasia mutant,
jun kinase, and mitogen-activated protein kinases.
The C-terminus undergoes phosphorylation by casein
kinase II, various cyclin-dependent kinases, and pro-
tein kinase C. Acetylation sites that also contribute
to the activation process have recently been identi-
fied (reviewed in [4,7,12]). Phosphorylation of the
N-terminus sets in motion a conformational change
that enables transcription to proceed, has a minor
effect on DNA binding, and contributes to the stabi-
lization of p53 by decreasing the interaction of p53
with mdm2 (reviewed in [6]). Changes in the phos-
phorylation of the carboxy terminus alter the con-
formation of the protein to modulate DNA binding
activity (reviewed in [4]). For example, dephospho-
rylation of serine 376 creates a binding site for the
14-3-3s protein, which when bound to p53, increases
the binding capacity of p53 [13]. In addition to phos-
phorylation and dephosphorylation, the C-terminus
undergoes acetylation at lysine 320 and 382 [14] and
binding to the small ubiquitin-like protein, SUMO-1,
at lysine 386 [15,16] as part of the activation process.
Because mutations in the p53 gene are so common
in human tumors and have been widely reported in the
literature, extensive databases exist such as that main-
tained by IARC [17]. Walker et al. [18] have utilized
the IARC database to define 73 “hotspots” for muta-
tion in p53 and related the mutations to changes in
protein structure and function. Most “hotspots” were
found at CpG dinucleotides within exons 5–8 [18].
Methylation of CpG sites lends itself to the induction
of mutation due to the tendency for deamination of
cytosine to uracil and the insertion of an inappropri-
ate thymine during repair (reviewed in [19,20]). It has
also been suggested that the binding of certain carcino-
gens is targeted to methylated CpG sites (reviewed in
[20]). Although the mechanism is not fully defined,
six CpG “hotspots” have been identified within the
DNA binding domain of human p53 and are found at
codons 175, 213, 245, 248, 273 and 282. An additional
“hotspot” has been found at codon 249 in aflatoxin
B1-induced hepatocellular carcinoma [21]. The effect
of these mutations on the structure of p53 has been ex-
amined by Cho et al. [22] utilizing X-ray crystallogra-
phy and provides convincing data that these mutations
alter the ability of p53 to bind to the promoter regions
of genes under its regulatory influence. Additional
“hotspots” have been identified outside of the DNA
![Page 3: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/3.jpg)
S.M. Morris /Mutation Research 511 (2002) 45–62 47
binding domain, but the mutation spectrum is much
broader than that observed in exons 5–8. Nonsense,
frame-shifts and a small proportion of missense mu-
tations have been found outside of exons 5–8 [18,23].
3. Function of the p53 protein
Models have been proposed in which p53 functions
directly in DNA replication, repair and recombination
in order to eliminate spontaneous and chemically-
induced DNA damage (see Fig. 1). Once damage
occurs, p53 undergoes phosphorylation, dephospho-
rylation, acetylation and sumoylation at specific sites
in order to serve as a transcription factor. In this role,
p53 regulates the synthesis of proteins that participate
in cellular functions important in the elimination of
cells with DNA damage or preventing the replication
of damaged cells [7,24,25].
The association between loss of p53 function and
malignant progression is partially due to the role p53
plays in the response to DNA damage as “guardian
of the genome”. Initial studies on the cell-cycle with
ionizing radiation (IR)-exposed cells led to the finding
that the G1 checkpoint is absent in cells with mutant
p53 [26]. Subsequently, it was found that p21waf−1,
an integral component of the G1 checkpoint, is under
the transcriptional control of p53 and that synthesis
of p21 increases after exposure to either IR or chem-
ical mutagens in cells with wild-type p53 [27]. A G2
checkpoint is intact in many cell lines with mutations
in p53 [28], but evidence exists that a p53-regulated
checkpoint also operates during the G2 transition and
that loss of this checkpoint contributes to an increase
in polyploid cells [29].
Apoptosis or programmed cell death is an ordered
event under genetic control [30] with multiple path-
ways leading to the final destruction of the cell. The
p53 protein is an integral component of one apopto-
sis pathway that responds to DNA damage such as
IR, topoisomerase inhibitors, and mutagens that form
covalent DNA adducts. P53 responds to DNA damage
signals, which may be the blockage of RNA poly-
merase II [31], by upregulating the synthesis of bax,
an apoptosis-promoting protein. The p53 protein also
down-regulates bcl-2, an apoptosis-inhibitory protein
[32–34], by interacting with the TATA binding pro-
moter to repress bcl-2 expression [35]. The Bax and
bcl-2 form homo- and heterodimers and function in the
control of the mitochondrial permeability transition
and the progression to the execution phase of apop-
tosis (reviewed in [36]). Interestingly, recent studies
have demonstrated that p53 translocates to the mito-
chondria and is sensitive to the levels of bcl-2 and bax
in the mitochondria [37,38]. A feedback loop for p53
and bcl-2 has recently been described in which bcl-2
inhibits the transactivation of the p53-regulated pro-
teins bax, p21waf−1, mdm2, cyclin G, and GADD45
promoters [39,40]. The p53 also functions as a tran-
scriptional regulator in the mitochondrial-independent
forms of apoptosis in which signals converge from
death receptors such as FAS/APO-1, killer/DR5, and
DR4 and the p53-inducible genes, or PIG’s which
respond to oxidative damage at the level of caspase-3
(reviewed in [41,42]).
Evidence is accumulating that p53 plays an
important role in DNA excision repair, both as a
transcription factor and as a component of nucleotide
and base excision repair complexes. It has recently
been shown that p53 functions in base excision repair
[43], to upregulate the synthesis of the repair enzyme,
O6-alkylguanine transferase [44], and to upregulate
the p48 subunit of the DNA damage binding protein
2 (DDBP2) which is involved in the binding of the
“global genomic repair” complex to DNA damaged
by UV [45]. Loss of p53 function in UV-exposed
Li–Fraumeni p53 mutant cells resulted in a decrease
in global genomic repair, but did not affect the rate
or efficiency of transcription-coupled repair [46,47].
Transfection of wild-type p53 human fibroblasts with
human papillomavirus Type 16 open reading frame
E6 (HPV16E6) and the subsequent loss of wild-type
p53 activity also resulted in the loss of global genomic
repair [48–50]. In addition to regulating the synthesis
of DBBP2 or p48, p53 physically interacts with XPB
and XPD, components of the nucleotide excision
repair (NER)-associated TFIIH transcription/repair
complex with helicase activity [51]. Through its
C-terminus, p53 binds to single-stranded DNA ends
[52] and participates in strand exchange activity
[53,54]. Sequence-specific binding by the C-terminus
of p53 is postulated to be a step in damage recog-
nition as part of its repair function [54]. However,
Liu and Kulesz-Martin [55] have proposed that the
DNA binding function of p53 may serve as a sensor
for directing damaged cells to the repair pathways or
![Page 4: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/4.jpg)
48 S.M. Morris /Mutation Research 511 (2002) 45–62
Fig. 1. Normal p53 response to DNA damage. Cellular DNA is damaged by chemical or physical insult (1); p53 is upregulated (2)
and undergoes phosphorylation, dephosphorylation and acetylation (3) to active isoforms; p53 acts as a transcription factor (4a) and as
a structural component (4b) of protein complexes; the proteins under the regulatory influence of p53 and the p53 protein complexes all
modulate cellular functions that influence the mutant frequency (5); cells are halted at the various p53-regulated checkpoints (6) to allow
the removal (repair) of DNA damage (7); when DNA damage is not repaired, the cells are then targeted for death (8); these factors
combine to reduce the accumulation of mutations within the genome (9). When mutation in p53 leads either to the loss of the protein or to
aberrant function of the protein, the p53-mediated response to DNA damage is abrogated, leading to an increase in the mutant frequency.
![Page 5: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/5.jpg)
S.M. Morris /Mutation Research 511 (2002) 45–62 49
toward the apoptosis pathways, rather than serve a
direct repair function.
The recombinational activity of p53 is separate from
its transactivational properties [24,56] and mutations
in p53 increase the rate of recombination [24,56–58].
The p53 protein interacts with the proteins, RAD51,
topoisomerase-I (topo-I) and topoisomerase-II
(topo-II)a, each of which plays a role in recombi-
nation, DNA repair and DNA replication. The p53
protein interacts with RAD51 through two binding
sites on p53 (amino acids 94–160 and amino acids
264–315). Binding was reduced, but not eliminated,
when the binding efficiency of p53 with specific mu-
tations (135Y, 249S, and 273H) was compared to that
of wild-type p53. The presence of a mutation in p53
resulted in a decrease in the efficiency of binding to
RAD51 [59]. Wild-type p53 recognizes three-stranded
DNA junctions that structurally resemble early re-
combination intermediates [60]. Further, the 3′→ 5′
exonuclease activity of p53 [61] appears to target the
destruction of these intermediates [62]. The p53 mu-
tants, especially the 273H mutant, are defective in this
ability [24] contributing to the lack of recombinational
control in certain p53 mutants [24]. A more indirect
role for p53 in recombination involves its transcrip-
tional control of bcl-2. In a recent study with exciting
implications for the understanding of recombination,
Saintigny et al. [63] have demonstrated that bcl-2 in-
hibits RAD51-mediated conservative, or homologous,
recombination. RAD51 undergoes post-translational
modification by bcl-2 which then shifts the processing
of double strand breaks to an error-prone pathway and
produces an increase in the mutant frequency. Further,
the effects of bcl-2 on recombination are separate and
distinct from its effects on the cell-cycle and apopto-
sis. These factors may contribute to the genomic insta-
bility observed in p53 mutant cells (reviewed in [64]).
The interaction of p53 with topo-I is mediated
through a binding site that is localized to the carboxy
terminus of p53 [65–67]. The p53/topo-I complex is
found at the site of the “cleavable complex” which is
the site of the breakage and religation that occurs as
part of topo-I activity [68]. The 273H mutation trans-
fected into HT 29 cells and the 239S, 245S and 273H
mutations transfected into Sf-9 insect cells all result
in a constitutive, rather than transient, association be-
tween p53 and topo-I [65,66] and possibly contribut-
ing to the stabilization of the “cleavable complex”.
The observations that both p53 and topo-II are
involved in the DNA breakage–ligation mechanisms
associated with recombination and replication led
Wang et al. [69] to study the relationship between the
two proteins. The p53 protein was found to serve as
a transcriptional repressor of topo-II, reducing syn-
thesis through an interaction with an ICE element
in the topo-II promoter. Neither a mutation at codon
175 nor the double mutation at codons 22 and 23
repressed the synthesis of topo-II. This may account
for the overexpression of topo-II observed in hepato-
cellular carcinoma [70]. However, Yuwen et al. [70]
also described an interaction between p53 and topo-II
that was detected by co-immunoprecipitation which
was later confirmed by others [71,72]. The interaction
occurs via the C-terminal region of p53 and similar
binding efficiencies were found for wild-type p53 and
mutant p53 (M237R, M237I and R273C) [72].
4. Mechanisms for loss-of-function
of wild-type p53
The p53 protein exists primarily as a tetramer,
forming the complex through the oligomerization
domain at the C-terminus of the protein [73]. When
mutant and wild-type subunits form a heterotetramer,
the mutant subunit drives the tetramer into the mu-
tant conformation in a dominant-negative manner.
Loss-of-function that is associated with heterozy-
gosity in the p53 gene is thought to be due to this
proposed dominant-negative mode of action [74,75].
The role of base-pair substitution mutations on
dominant-negative interactions was evaluated utilizing
wild-type p53 colon adenocarcinoma cells transfected
with plasmids carrying different point mutations in
p53 [76]. Loss-of-function, as determined by the lack
of mdm2 synthesis and G1 arrest after IR exposure
and dominant-negative interaction, were dependent
upon mutation that resulted in a sequence change
in the protein [76]. Further, the mutant protein may
interfere with normal cellular processes and possess
a “gain-of-function” phenotype (reviewed in [77]).
Recently, p53 has been described as a member
of a “supergene” family with members p63 and
p73 exhibiting structural and functional similarity
to p53. In contrast to p53, however, both p63 and
p73 encode multiple splicing variants that encompass
![Page 6: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/6.jpg)
50 S.M. Morris /Mutation Research 511 (2002) 45–62
the C-terminal region (reviewed in [78]). P73,
described independently by Jost et al. [79] and by
Kaghad et al. [80], is characterized by the presence
of DNA binding domain CpG “hot spots” for muta-
tion, contains 14 exons and acts a transcription factor
for several p53-regulated genes, including those that
regulate apoptosis [81]. The p73 protein responds to
DNA damage by undergoing post-translational mod-
ification by the tyrosine kinase activity of c-abl and
inducing the apoptotic cascade [82–84]. In one study
[85], it was suggested that neither wild-type p53 nor
mutant p53 (R273H) formed heteroligomers with
p73. DiComo et al. [86], however, found that while
wild-type p53 did not oligomerize with wild-type
p73, mutant p53 (R175H and 248W) was found to
co-immunoprecipitate with p73, indicative of an inter-
action. Marin et al. [87], however, have demonstrated
that the strength of the interaction between certain
mutant forms of p53 and p73 may be determined by
the presence of a polymorphism at codon 72 of p53.
This polymorphism does not appear to affect p53–p53
interaction, but does demonstrate that certain muta-
tions in p53 proteins can lead to an interaction with
p73. P63 is composed of 15 exons and bears amino
acid homology to p53 in the transactivation domain,
DNA binding domain, and oligomerization domain
(reviewed in [78]). Neither the full length nor the
truncated splicing variants of p63 have been shown
to interact with p53 in a dominant-negative manner
[88].
Nuclear and cytoplasmic p53 levels are modulated
in part by the p53/mdm2/ARF pathway which serves
as a point of convergence for oncogenic signals such
as ras, E2F1, and c-myc (reviewed in [89,90]). The
p53 protein undergoes ubiquitination by the E3 ligase
activity of mdm2 and is shuttled from the nucleus to
the cytoplasm for degradation by the 26S proteosome
(reviewed in [91]). Mutation in p53 that results in
conformational alteration may inhibit mdm2’s ability
to bind p53 and target the protein for destruction,
thus, leading to increased nuclear levels of the mutant
protein [91]. The mdm2 ligase activity is under the
control of the alternate reading frame (ARF) proteins,
p14ARF (human) and p19ARF (mouse) which are
encoded in the INK4A/ARF locus [92]. In addition
to ligase activity, ARFs are also involved in the se-
questration of mdm2 in the nucleolus [93,94]. The
ARF-mediated decrease in mdm2 levels is paralleled
by an increase in the nuclear p53 levels which allows
the p53-directed transcription of the genes important
in cell proliferation, apoptosis, and perhaps, DNA
repair, to proceed. Mutation in the INK4A/ARF lo-
cus is quite common in malignancies and the loss of
ARF-mediated control of mdm2 activity can result in
biological responses similar to those observed in cells
with mutation in p53 (reviewed in [95]).
Another mechanism for the loss of wild-type func-
tion is the interaction with viral proteins. The SV40
protein, whose interaction with p53 led to its dis-
covery, binds to the DNA binding domain of p53
eliminating its functional ability [96]. HPV16E6 on-
coprotein binds to the p53 protein and transports it to
the ubiquitin pathway resulting in premature degra-
dation and a functionally p53 “null” state in the cell
[97]. A recent study also demonstrates that HPV16E6
down-regulates p53 by binding to CBP/p300 [98,99].
The adenovirus E1B-55KD protein abrogates p53
function, possibly by interacting with the RNA poly-
merase II transcription complex [100,101].
5. Cell line families that differ in p53 status
One of the original studies to suggest a role for p53
in modulating the mutant frequency was conducted in
TK6 human lymphoblastoid cells ([102]; see Tables 1
and 2). TK6 was derived from the human lymphoblas-
toid cell line, WIL2, as was the “closely-related” cell
line, WTK1 [102]. Both lines are heterozygous for the
thymidine kinase (TK) gene with frame-shift muta-
tions in exon 4 of the mutant TK allele and exon 7 of
the wild-type allele [103]. IR produces a much greater
frequency of TK mutants in WTK1 cells than in TK6
cells [102]. Molecular analysis of the TK gene in mu-
tant clones [104] revealed that the difference in mutant
frequency was due to an increase in the frequency of
a specific class of mutant recovered in WTK1 cells.
Loss-of-heterozygosity (LOH), due either to a deletion
mechanism or to errors in recombinational repair, was
detected at a much higher frequency in WTK1. Subse-
quent studies revealed that the two lines differed in p53
status with a homozygous G → A mutation present at
codon 237 of exon 7 ofWTK1 (andWIL2-NS, another
TK6-derived cell line) [105,106]. The codon 237 mu-
tation lies in the DNA binding region of p53 and the
homozygous mutation would be expected to interfere
![Page 7: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/7.jpg)
S.M. Morris /Mutation Research 511 (2002) 45–62 51
Table 1
Cell lines, their p53 status and reporter genes used for mutant frequency evaluationa
Parent cell line Cell line p53 status Loss of p53 function due to Reporter gene
WIL2 (H) TK6 Wild-type p53 function intact TK, HPRT
TK6-E6 Wild-type Targeted degradation of p53 by HPV16E6 TK, HPRT
TK6-NH32 Mutant (null) Homozygous deletion TK, HPRT
WTK1 Mutant (point) Homozygous C → T at codon 237 (exon 7) TK, HPRT
AHH-1 (H) L3 Wild-type p53 function intact TK, HPRT
MCL-5 Wild-type p53 function intact TK, HPRT
AHH-1 TK+/− Mutant (point) Heterozygous C → T at codon 282 (exon 8) TK, HPRT
SAOS-2 (H) SAOS-2 Mutant (null) Homozygous deletion of exons 2–11 HPRT
RKO (H) RKO Wild-type p53 function intact HPRT
RKO-E6 Wild-type Targeted degradation of p53 by HPV16E6 HPRT
LN12 (M) LN12 Mutant Deletion supF
a H: human origin; M: mouse origin.
with the transcription factor properties associated with
p53 as well as affecting p53-mediated recombination.
In a series of studies, Honma et al. [107,108] have
elegantly demonstrated that although spontaneous and
IR-induced TK mutations in both TK6 and WTK1
cells resulted from LOH, the mechanism leading to
the mutation differed as a function of p53 status. TK
mutants were identified as LOH by Southern blot pat-
terns and then classified as homozygous (two copies
of the non-functional allele) or hemizygous (one copy
of the non-functional allele) by densitometric analysis.
In TK6 cells, both hemizygous and homozygous LOH
was detected, in contrast to WTK1 cells, in which
two non-functional alleles were detected in the major-
ity of clones [107]. When representative clones were
analyzed by chromosome painting, it was revealed
that the LOH in TK6 cells could not be accounted
for by chromosome rearrangements. In WTK1 cells,
however, a high percentage of translocations was
found in both the spontaneous and IR-induced clones
[108]. These findings led to a model in which the
cells with mutant p53, WTK1, had a predisposition
towards non-homologous recombination that could be
followed by mitotic non-disjunction. In contrast, ho-
mologous recombination predominated in TK6 cells
with wild-type p53 [108].
When the same mutation in p53 found in WTK1
cells, H237I, was transfected into wild-type TK6
cells, both the spontaneous and the IR-induced mutant
frequencies increased to the level observed in WTK1
cells [109]. These findings led to the question of
whether the increase in the mutant frequency was due
to the presence of the mutant protein or to the absence
of the wild-type protein. Thus, TK6 cells were trans-
fected with high-risk, HPV16E6, effectively creating a
p53 “null” cell line (TK6-E6), due to the rapid degra-
dation of the p53 protein. The spontaneous mutant fre-
quency in TK6-E6 cells was elevated in comparison to
TK6 cells, but not to the degree observed in the WTK1
line [110,111]. Exposure of TK-E6 cells to IR resulted
in a TK mutant frequency intermediate to IR-exposed
TK6 and WTK1 cells [111]. Molecular and cytoge-
netic analyses were performed on spontaneous mu-
tants recovered from the TK6, the TK6-E6, and the
vector control, TK6-20C lines [110]. Quantitative PCR
revealed the loss of the functional TK allele in mutants
derived from each of the cell lines. The mutations re-
covered from TK6-E6 cells in which p53 underwent
viral degradation were predominately hemizygous
and the loss of the terminal portion of chromosome
17q was confirmed by microsatellite analysis. Sub-
sequent chromosome painting analysis revealed that
allele loss and the formation of deletions and translo-
cations could be attributed to errors in end-rejoining
and non-homologous recombination. In contrast, both
hemizygous and homozygous TK mutants were de-
rived from both the TK6 and TK6-20C cell lines in
which p53 remains functionally intact [110].
A confounding factor in the experiments that uti-
lize TK6-E6 cells is the observation that the effects
of HPV16E6 may not be limited to the destruction
of p53 and that those effects may contribute to the
![Page 8: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/8.jpg)
52 S.M. Morris /Mutation Research 511 (2002) 45–62
![Page 9: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/9.jpg)
S.M. Morris /Mutation Research 511 (2002) 45–62 53
differences in the TK mutant frequency. To address
this question, Chuang et al. [112] developed a TK6
derivative, TK6-NH32, in which both alleles of p53
were disrupted by molecular targeting resulting in a
p53 “null” genotype with neither wild-type nor mu-
tant p53. When cells from the TK6, TK6-NH32, and
WTK1 cell lines were exposed to IR and the spon-
taneous and IR-induced TK mutant frequencies mea-
sured, the increase in the mutant frequency in cultures
derived from WTK1 was substantially greater than
that observed in either TK6 or TK6-NH32 cells. These
data led to the suggestion that the mutation in p53 in
the WTK1 line led to a “gain-of-function” which con-
tributed to the mutant frequency to a greater extent
than the “loss-of-function” mutation in the TK6-NH32
line [112].
Evidence indicating a role for p53 in determining
the mutant frequency also has come from studies
with the AHH-1 cell line family. AHH-1 TK+/− and
MCL-5 (L3) cells were derived from the AHH-1 cell
line described by Crespi and Thilly [113]. Both are
heterozygous for the TK gene with a frame-shift mu-
tation in exon 4 of the mutant allele [114]. Sequence
analysis of the p53 gene in these cells revealed that
a C → T transition had occurred at codon 282 of
exon 8 of AHH-1 TK+/− cells. This is a reported
“hot spot” for C → T mutations and results in an
amino acid substitution in the DNA binding domain
of the protein [115]. Several classes of compounds
produce a five-fold greater TK mutant frequency in
AHH-1 TK+/− cells than in MCL-5 (L3) cells. When
AHH-1 TK+/− and L3 cells were exposed to the pu-
tative topo-II inhibitors and phytoestrogens, genistein
and coumestrol [116,117], a five-fold increase in the
percentage of TK mutant clones with the slow-growth
phenotype was found. When a screening assay for
LOH was utilized to evaluate the coumestrol-induced
TK mutant clones, wild-type exon 4 was not detected.
Mutagen-induced LOH was also detected in the ex-
periments of Dobo et al. [118] in which a higher
frequency of TK mutants was found in the AHH-1
TK+/− cell line. Deletion analysis by microsatellite
loci revealed that the length of the LOH tracts were
substantially longer in the AHH-1 TK+/− cell line.
The increased mutant frequency in the TK gene of
AHH-1 TK+/− cells was accompanied by a decreased
rate of apoptosis and the loss of the G1 checkpoint,
factors consistent with the mutation in p53 [116,118].
Studies addressing the role of p53 on mutant fre-
quency have not been limited to the TK gene. Sev-
eral studies have examined the effect of p53 status
on hypoxanthine-guanine phosphoribosyl transferase
(HPRT) mutant frequency and on the frequency of
mutations in the bacterial reporter gene, supF, which
has been integrated into mouse LN12 cells. Wild-type
p53, under the control of an inducible promoter, was
transfected into SAOS-2 cells which are derived from
a human osteosarcoma and deficient in p53 expression
due to a deletion encompassing exons 2–11 [119].
HPRT mutant frequencies, measured after exposure to
X-rays [120] or to UV [121], were markedly reduced
in the clones that expressed wild-type p53 compared
to the p53 null cells. UV exposure of RKO cells trans-
fected with HPV16E6 resulted in mutant frequencies
in the HPRT gene greater than non-transfected RKO
cells [122]. A similar approach, the transfection and
expression of wild-type p53 into p53-deficient cells,
was utilized by Yuan et al. [123] in mouse LN12
cells. Mutations in the supF reporter gene decreased
four-fold in the wild-type p53-expressing cell line.
Molecular analysis was not performed in the exper-
iments with SAOS-2 or RKO cells. Finally, IR pro-
duced a higher mutant frequency in the HPRT gene
in WTK1 cells than in TK6 cells [124]. However, no
differences in the mutation spectra were detected.
6. In vivo mutation detection systems
Although cell culture models have increased our
knowledge of p53 function in maintaining genomic in-
tegrity, understanding the p53-modulated response to
DNA damaging agents in an animal or human model
involves a greater level of complexity than in a cell
culture system. A major advance in modeling the role
of p53 has been the production of three strains of trans-
genic mice in which a major deletion in p53 was in-
troduced by recombinant DNA technology [125–127].
Differences among the strains exist in that intron 4 and
exon 5 were deleted in the Donehower strain [125] in
contrast to the deletion of exons 2 through 6 in the
models developed by Jacks et al. [126] and by Purdie
et al. [127]. In addition, the genetic background differs
among the strains with both the Donehower and Jacks
strains being derived on a C57Bl/6 × 129/SV back-
ground and the Purdie strain on a 129/Ola background.
![Page 10: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/10.jpg)
54 S.M. Morris /Mutation Research 511 (2002) 45–62
However, common to each of these strains is a propen-
sity to develop thymic lymphoma at an early age.
The tumor spectrum of the p53 heterozygous mouse
differs from that found in the null mouse in that there
is a predisposition towards the induction of sarcomas
rather than lymphoma. As discussed in Purdie et al.
[127] and MacLeod and Jacks [128], the difference
in the spectrum between the “knockout” and the het-
erozygote may reflect the susceptibility of the tissue
for the loss of the second p53 allele. Differences in the
characteristics of the heterozygotes exist between the
strains. In the Donehower strain [125], p53 levels are
reduced below what would be expected on the basis of
allele dosage and the lowered level of p53 may con-
tribute to the phenotype of this p53+/− mouse [129].
Table 3
The p53 status, reporter locus, and mutant frequency (MF) in transgenic mice
Reporter locus Tissue Mutagen Mutant frequency Reference
LacI Liver None p53+/+= p53−/− [130]
Spleen p53+/+= p53−/−
Brain p53+/+= p53−/−
LacI Thymic
lymphoma
None Increase of 3× in one of the four tumors (2.3 × 10−5 in p53+/+
mice vs. 6.8 × 10−5 in one tumor from p53−/− mouse)
[132]
LacI Embryonic
fibroblast
2mM 4-NQO No p53-mediated difference in spontaneous MF; no
p53-mediated difference in MF after exposure to 2mM 4-NQO
although the LacI MF increased in response to 4-NQO exposure
[133]
Thymus None p53+/+= p53−/−
Thymic
lymphoma
None p53+/+= p53−/−
APRT Fibroblasts None Increase of 3× in spontaneous MF (10−5 for p53+/− mice and
33.5 × 10−5 for p53−/− mice); molecular analysis revealed
mutation due to mitotic recombination which results in LOH
[136]
T-lymphocytes p53+/−= p53−/−
pun Mouse pups B[a]P 30mg/kg Control litters were 12–16% “spotted” or mutant pups; 40–50%
of the pups from exposed dams of each genotype were “spotted”
[140]
B[a]P 150mg/kg Control litters were 12–16% “spotted” pups; 77–100% of the
pups from exposed dams were “spotted”
IR 1Gy Reduced litter sizes in all genotypes, but most predominately in
the IR-exposed p53−/− litters. No “spotted” pups were present
at 10 days post-partum in p53−/− litters
LacZ Lung 13mg/kg of B[a]P
for 13 weeks
p53+/+= p53+/− [142]
Liver p53+/+= p53+/−
Spleen p53+/+= p53+/−
LacZ Spleen 300 ppm of AAF
for 12 weeks
p53+/+= p53+/− [143]
Liver p53+/+= p53+/−
Bladder p53+/+ < p53+/−
Many of the tumors in the Donehower heterozygote re-
tain the wild-type allele and tumor induction is linked
to the lowered gene dosage [129]. P53 levels are at ap-
proximately 50% of that found in the wild-type mouse
in the Jacks strain [126]. Further, LOH of the p53 gene
occurs in a high proportion of the tumors found in the
Jacks strain apparently by a chromosomal mechanism.
Each of the p53 transgenic mouse models has been
utilized to explore the effect of mutations in the p53
gene on the frequency and spectrum of mutations at
reporter genes (see Table 3). Utilizing the Donehower
model [125], animals of each of the p53 genotypes
(p53+/+; p53+/−; p53−/−) and bearing the LacI
transgene (TSG-p53/Big Blue©) were created. Spon-
taneous mutant frequencies were measured in liver,
![Page 11: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/11.jpg)
S.M. Morris /Mutation Research 511 (2002) 45–62 55
spleen and brain [130,131], and after correcting for
clonal expansion of single mutations, no difference
in the LacI mutant frequency as a function of p53
genotype was observed. The spontaneous LacI mutant
frequency was also determined in thymic lymphomas
derived from p53−/−/LacI+/− mice. A 2.3-fold in-
crease in the mutant frequency (6.8 × 10−5 versus
2.9 × 10−5 in p53−/− thymus cells) was detected in
one of four tumors with an increase in A : T → G : C
transition mutations. Two tumors demonstrated a
trend towards an increase in the mutant frequency
(3.7 × 10−5 versus 2.9 × 10−5 in p53−/− thymus
cells) and no increase in the mutant frequency was
found in the other tumor [132].
The p53+/−/LacI mouse has also been utilized
to examine the relationship between p53 status
and both spontaneous and 4-nitroquinoline oxide
(4-NQO)-induced mutations. The spontaneous LacI
mutant frequencies were measured in embryonic
fibroblasts, thymocytes derived from 2–3 months
old animals and thymomas detected in animals 5–8
months old. In addition, embryonic fibroblasts derived
from p53+/+/LacI and p53−/−/LacI animals were ex-
posed to 2mM 4-NQO. Although a dose-responsive
increase in the LacI mutant frequency was found in
the 4-NQO-treated fibroblasts, no effect of p53 status
was found with any of the three genotypes [133].
Studies with the LacI reporter gene may be limited
since the LacI is a transgene, rather than an endoge-
nous gene, and does not efficiently detect chromo-
some damage and recombination events [130]. In
contrast, adenine phosphoribosyl transferase (APRT)
is an endogenous, autosomal gene, capable of de-
tecting these types of events. A mouse model, de-
veloped by Stambrook and co-workers [134,135],
is heterozygous for the APRT gene and responds to
mutagen exposure with an increase in APRT mutant
frequency. Thus, the APRT+/− mouse was crossed
with p53+/− mice to create animals with the geno-
types of p53+/+/APRT+/−, p53+/−/APRT+/−, and
p53−/−/APRT+/−. Mutant frequencies in the APRT
gene, measured by resistance to 2,6-diaminopurine
(DAP) were similar in the T-lymphocytes of the
p53+/+ and p53−/− mice. However, a three-fold
increase in the frequency of DAP-resistant variants
derived from fibroblasts was found in p53−/− mice
compared to p53+/− mice. The APRT mutant fre-
quency in the fibroblasts of p53+/− mice did not differ
significantly from that of the p53+/+ animals. Mitotic
recombination accounted for the majority of mutations
in the wild-type, the heterozygous and the “knockout”
mice; however, deletion and gene conversion ac-
counted for 5 out of 47 variants in the p53+/− mice
and 6 of 106 variants in the p53−/− mice. An increased
number of variants (7/106) resulting from chromo-
some damage (non-homologous recombination) were
encountered in the p53−/− fibroblast clones [136].
Another locus which has been utilized to examine
the effect of p53 status on mutant frequency is the
pink-eyed dilution unstable (pun) mutation which is
maintained in the C57Bl/6J mouse strain. This mu-
tation results from the tandem duplication of 70 kb
internal to the “p” gene that is expressed in embryonic
melanocytes [137,138]. Reversion to wild-type occurs
by the deletion of one of the duplicated sequences
and can be measured phenotypically by the presence
of black spots on the grey coat of the mouse. The
reversion frequency of the pun mutation in mice with
wild-type p53 increases with exposure to IR [139].
When mice of each of the p53 genotypes were crossed
with the pun mouse and pregnant dams exposed to IR,
an increase in the number of “spotted” offspring was
found in the p53+/+ and p53+/− offspring, but not
in the p53−/− pups. It was suggested that IR-induced
damage was processed differently in the p53−/− pups
and that this could account for the lack of “spotted”
or mutant pups [140]. When pregnant dams were ex-
posed to the point mutagen, benzo(a)pyrene (B[a]P),
p53 status did not affect the reversion frequency [140].
NER-defective mice, Xpa [141], were crossed with
the Jacks strain [126] to create a transgenic mouse with
the Xpa−/−/p53+/− genotype. In order to study the ef-
fect of the loss of NER and p53 heterozygosity on mu-
tant frequency, a triple transgenic mouse was created
by crossing this animal to a strain harboring a plasmid
vector containing the LacZ transgene [142,143]. When
exposed to B[a]P, the LacZ mutant frequencies in the
lungs, liver and spleens of the p53+/− mice were not
significantly elevated compared to those observed in
the p53+/+ animals. However, when exposed to the
carcinogen, 2-acetylaminofluorene (2-AAF), the LacZ
mutant frequency in the bladder, but not the spleen
or liver, increased significantly in the p53+/− animals
compared to the wild-type. Thus, the effect of p53 het-
erozgosity on LacZ mutation induction appears to be
restricted to specific tissues [143].
![Page 12: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/12.jpg)
56 S.M. Morris /Mutation Research 511 (2002) 45–62
The strains developed by Jacks et al. [126] were uti-
lized to examine the effect of IR on the HPRT mutant
frequency in preB cells that are sensitive to IR [144].
Although IR-induced 6-thioguanine-resistant clones
were recovered from p53−/− mice, none was recov-
ered from the wild-type mice due to an extremely low
cloning efficiency. Southern analysis confirmed the
loss of multiple exons within the HPRT gene.
7. Summary
Clear and consistent evidence is emerging that
mutations in p53 affect both the frequency and the
pathway leading to mutation. In models in which loss
of p53 function occurs due to point mutation, targeted
deletion, or loss of protein due to viral degradation,
there is a consistent increase in the TK mutant fre-
quency, and the increase is due, in most instances, to
mechanisms that result in LOH. Loss of p53 func-
tion leads to an increase in the frequency of mutation
due to non-homologous recombination that may be
followed by mitotic non-disjunction and result in the
formation of chromosome rearrangements. In cells
with wild-type p53, mutation that results in LOH
seems to occur preferentially through a homologous
recombination pathway that results in the loss of the
wild-type allele.
Further, the type of mutation in p53 may play a
major role in determining the mutant frequency in
the reporter gene. Most point mutations in p53 that
have been extensively studied are those that reside
in the DNA binding domain and in the classic CpG
“hotspots”. Studies with the representative DNA
binding domain mutant, 273H, indicate that recombi-
nation activities are hindered by the altered binding
of mutant p53 to recombination pathway proteins
such as RAD51. The down-regulation of bcl-2 by p53
would be reduced in p53 mutant cells, resulting in an
increase in bcl-2-induced modification of RAD51 and
a shift to error-prone pathways. Thus, recombination
in cells with mutant p53 would more likely result
in damage being processed through “error-prone”
pathways and a high mutant frequency. In addition,
mutant p53 protein interferes with other cellular
processes that may affect the mutant frequency, in-
cluding apoptosis [145]. The importance of apoptosis
in determining mutant frequency is suggested by
experiments in which a decreased rate of apoptosis is
accompanied by an increase in TK mutant frequency
[116,118]. Additional support for this hypothesis
comes from the studies of Cherbonnel-Lassere et al.
[146,147] in which ectopic overexpression of the
p53-regulated apoptosis-inhibitory protein, bcl-2, re-
sulted in an increase in spontaneous and induced TK
mutant frequency. Although the increased expression
of bcl-2 may result in a shift to error-prone pathways,
apoptosis is also inhibited by high levels of bcl-2
and is manifested in an increase in clonogenic sur-
vival. Cells that would normally be targeted for death
contribute to an increase in the mutant frequency.
A different pattern emerges in cells in which p53
function has been abrogated as a result of gene
deletion or viral infection. Recombination-mediated
mutations occur at a much lower frequency in p53
“null” cells, cells subjected to viral infection, and
wild-type cells than in p53 mutant cells. These results
are consistent with the hypothesis that mutant protein
interferes with the pathways involved in the repair
and removal of cells with DNA damage. The lack of
wild-type p53 protein may affect the recombination
processes, perhaps through monitoring and degrading
early recombination intermediates, or may shift the
DNA damage response to an alternate pathway, such
as apoptosis. Mutant p53 protein may interfere with
recombination, apoptosis, and other cellular processes
and contribute to the high levels of mutations resulting
in LOH. The absence of mutant protein may possibly
account for the minimal increase in the mutant fre-
quency not only in cell lines, but also in the in vivo
model systems that have a p53 “null” background. An
approach to testing this hypothesis would be to cross
a transgenic mouse that is heterozygous at a reporter
gene loci, e.g. the TK+/− mouse of Doborovolsky
et al. [148] or the APRT+/− mouse of Stambrook and
co-workers [134,135], with a mouse that carries a
point mutation in p53 such as that recently described
by Liu et al [149].
A body of evidence exists that mutation leading
to either the absence of the wild-type protein or the
presence of the mutant protein results in the decreased
ability of p53 to function both as a transcription fac-
tor and to properly serve as a structural component
of protein complexes involved in the response to
DNA damage. Experiments that more fully define the
mechanism(s) by which the presence of mutant p53
![Page 13: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/13.jpg)
S.M. Morris /Mutation Research 511 (2002) 45–62 57
results in a substantially greater increase in the mu-
tant frequency than does the absence of the wild-type
protein will be of considerable interest. An approach
which may provide insight into the effect of mutation
on the ability of p53 to serve as a transcription factor
would include genome-wide expression analysis of
cells with mutations in critical sites of the gene. The
effect of critical site mutations on the interaction of
p53 with other cellular proteins, as well as the iden-
tification of additional proteins that interact with p53,
could be characterized by protein “chip” analysis
combined with two-dimensional gel electrophoresis.
This knowledge would contribute substantially to our
understanding of the pathways leading to malignancy.
It is clear, however, that with the failure of the G1
checkpoint, the inability of cells to repair damaged
DNA before replication and the abrogation of the tar-
geted removal of cells with damaged DNA from the
population by apoptosis, damaged or mutant cells can
survive, proliferate and contribute to the progression
to malignancy. Further, p53 serves as a convergence
point for the integration of signals from oncogenic
stimuli as well as those from the DNA damage path-
ways and both pathways contribute to the cell’s abil-
ity to maintain genomic integrity. Through its ability
to modulate the acquisition of additional mutation,
p53 plays a central role in the guardianship of the
genome and loss of p53 contributes significantly to
the progression of cells to a malignant state.
Acknowledgements
The author would like to thank Dr. William Tolleson
and Dr. Greg Akerman for helpful discussions and Dr.
Robert Heflich for critical review of the manuscript.
References
[1] D.P. Lane, Cancer: p53, guardian of the genome, Nature
358 (1992) 15–16.
[2] X.W. Wang, C.C. Harris, Tp53 tumour suppressor gene:
clues to molecular carcinogenesis and cancer therapy, Cancer
Surveys, Genetic Instability in Cancer, Vol. 28, 1996,
pp. 169–196.
[3] L. Jayaraman, C. Prives, Covalent and noncovalent modifiers
of the p53 protein, Cell. Mol. Life Sci. 55 (1999) 76–87.
[4] W. Steegenga, A.J. van der Eb, A.G. Jochemsen, How
phosphorylation regulates the activity of p53, J. Mol. Biol.
263 (1996) 103–113.
[5] J.M. Milczarek, G.T. Bowden, p53 phosphorylation:
biochemical and functional consequences, Life Sci. 60
(1997) 1–11.
[6] W.D. El-Diery, The p53 pathway and cancer therapy, Cancer
J. 11 (1998) 229–236.
[7] D.W. Meek, New developments in the multi-site phospho-
rylation and integration of stress signaling at p53, Int. J.
Radiat. Biol. 74 (1998) 729–737.
[8] N. Albrectson, I. Dornreiter, F. Grosse, E. Kim, L.
Wiesmuller, W. Deppert, Maintenance of genomic integrity
by p53: complementary roles for activated and non-activated
p53, Oncogene 18 (1999) 7706–7717.
[9] C. Prives, P.A. Hall, The p53 pathway, J. Pathol. 187 (1999)
112–126.
[10] N.D. Lakin, S.P. Jackson, Regulation of p53 in response to
DNA damage, Oncogene 18 (1999) 7644–7655.
[11] G.S. Jimenez, S.H. Khan, J.M. Stommel, G.M. Wahl,
p53 regulation by translational modification and nuclear
retention in response to diverse stresses, Oncogene 18
(1999) 7656–7665.
[12] A. Apella, C.W. Anderson, Post-translational modifications
and activation of p53 by genotoxic stresses, Eur. J. Biochim.
268 (2001) 2764–2772.
[13] M.J.F. Waterman, J.L.F. Waterman, T.D. Halazonetis,
ATM-dependent activation of p53 involves dephosphoryla-
tion and association with 14-3-3 protein, Nat. Genet. 19
(1998) 175–178.
[14] W. Gu, R.G. Roeder, Activation of p53 sequence-specific
DNA binding by acetylation of the p53 C-terminal domain,
Cell 90 (1997) 595–606.
[15] M.S. Rodriguez, J.M.P. Desterro, S. Lain, C.A. Midgley,
D.P. Lane, R.T. Hay, SUMO-1 modification activates
the transcriptional response of p53, EMBO J. 18 (1999)
6455–6461.
[16] M. Gostissa, A. Hengerstermann, V. Fogal, P. Sandy, S.E.
Schwarz, M. Scheffner, G. Del Sal, Activation of p53 by
conjugation to the ubiquitin-like protein SUMO-1, EMBO
J. 18 (1999) 6462–6471.
[17] P. Hainut, T. Hernandez, A. Robinson, P. Rodriguez-Tome,
T. Flores, M. Hollstein, C.C. Harris, R. Montesano, IARC
database of p53 gene mutations in human tumors and
cell lines: updated compilation, revised formats and new
visualization tools, Nucl. Acids Res. 26 (1998) 205–213.
[18] D.R. Walker, J.P. Bond, R.E. Tarone, C.C. Harris, W.
Makalowski, M.S. Boguski, M.S. Greenblatt, Evolutionary
conservation and somatic mutation hotspots maps of p53:
correlation with p53 protein structural and functional
features, Oncogene 19 (1999) 211–218.
[19] M.S. Greenblatt, W.P. Bennet, M. Hollstein, C.C. Harris,
Mutations in the p53 tumor suppressor gene: clues to
cancer etiology and molecular pathogenesis, Cancer Res.
54 (1994) 4855–4878.
[20] G.P. Pfeifer, p53 mutational spectra and the role of methy-
lated CpG sequences (review), Mutat. Res. 450 (2000)
155–166.
[21] I.C. Hsu, R.A. Metcalf, T. Sun, J.A. Welsh, N.J. Wang,
C.C. Harris, Mutational hotspot in the p53 gene in human
hepatocellular carcinomas, Nature 350 (1991) 427–428.
![Page 14: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/14.jpg)
58 S.M. Morris /Mutation Research 511 (2002) 45–62
[22] Y. Cho, S. Gorina, P.D. Jeffrey, N.P. Pavletich, Crystal
structure of a p53 tumor suppressor–DNA complex: under-
standing tumorigenic mutations, Nature 265 (1994) 33346–
33355.
[23] X.W. Wang, C.C. Harris, p53 tumor-suppressor gene: clues
to molecular carcinogenesis, J. Cell. Phys. 173 (1997)
247–255.
[24] C. Dudenhoffer, M. Kurth, F. Janus, W. Deppert, L.
Weismuller, Dissociation of the recombination control
and the sequence-specific transactivation function of p53,
Oncogene 18 (1999) 5773–5784.
[25] F. Janus, N. Albrechtson, I. Dornreiter, L. Wiesmuller,
F. Grosse, W. Deppert, The dual role model for p53 in
maintaining genomic integrity, Cell. Mol. Life Sci. 55
(1999) 12–27.
[26] M.B. Kastan, O. Onywerke, D. Sidransky, B. Vogelstein,
R.W. Craig, Participation of p53 protein in the cellular res-
ponse to DNA damage, Cancer Res. 51 (1991) 6304–6311.
[27] W.S. El-Diery, T. Tokino, V.E. Velculisci, D.B. Levy, J.M.
Trent, D. Lin, W.E. Mercer, K.W. Kinzler, B. Vogelstein,
WAF1, a potential mediator of p53 tumor suppression, Cell
75 (1993) 817–825.
[28] S.M. Morris, L.J. McGarrity, O.E. Domon, J.J. Chen, D.A.
Casciano, Cell-cycle traverse in AHH-1 TK+/− human
lymphoblastoid cells exposed to the chromosomal mutagen,
m-amsa, Environ. Mol. Mutagen. 27 (1996) 10–18.
[29] S.A. Innocente, J.L.A. Abrahamson, J.F. Cogswell, J.M.
Lee, p53 regulates a G2 checkpoint through cyclin B1,
Proc. Natl. Acad. Sci. 96 (1999) 2147–2152.
[30] A.H. Wyllie, Apoptosis and the regulation of cell numbers
in normal and neoplastic tissues: an overview, Cancer
Metast. Rev. 11 (1992) 95–103.
[31] M. Ljungman, F. Zhang, F. Chen, A.J. Rainbow, B.C.
McKay, Inhibition of RNA polymerase II as a trigger for
the p53 response, Oncogene 18 (1999) 583–592.
[32] T. Miyashita, J.C. Reed, bcl-2 gene transfer increases
relative resistance of S49.1 and WEHI7.2 lymphoid cells to
cell death and DNA fragmentation induced glucocorticoids
and multiple chemotherapeutic drugs, Cancer Res. 52
(1992) 5407–5411.
[33] M. Selvakumaran, H.K. Lin, T. Miyashita, H.G. Wang,
S. Krajewski, J.C. Reed, B. Hoffman, D. Liebermann,
Immediate up-regulation of bax expression by p53 but not
by TGFb1: a paradigm for distinct apoptotic pathways,
Oncogene 9 (1994) 1791–1798.
[34] Q. Zhan, S. Fan, I. Bae, C. Guillouf, D.A. Libermann, P.M.
O’Conner, A.J. Fornace, Induction of bax by genotoxic
stress in human cells correlates with normal p53 status and
apoptosis, Oncogene 9 (1994) 3743–3751.
[35] Y.L. Wu, J.W. Mehew, C.A. Heckman, M. Arcinas, L.M.
Boxer, Negative regulation of bcl-2 expression by p53 in
hematopoietic cells, Oncogene 20 (2001) 240–251.
[36] M. Crompton, The mitochondrial permeability transition
pore and its role in cell death, Biochem. J. 341 (1999)
233–249.
[37] N.D. Marchenko, A. Zaika, U.M. Moll, Death signal-induced
localization of p53 protein to mitochondria: a potential
role in apoptotic signaling, J. Biol. Chem. 275 (2000)
16202–16212.
[38] R.J. Donahue, M. Razmara, J.B. Hoek, T.B. Knudsen, Direct
influence of the p53 tumor suppressor on mitochondrial
biogenesis and function, FASEB J. 15 (2001) 635–644.
[39] B.A. Froesch, C. Aime-Sempe, B. Leber, D. Andrews, J.C.
Reed, Inhibition of p53 transcriptional activity by bcl-2
requires its membrane-anchoring domain, J. Biol. Chem.
274 (1999) 6469–6475.
[40] Q. Zhan, U. Konty, M. Iglesias, I. Alamo, K. Yu, C.
Hollander, C.D. Woodsworth, A.J. Fornace, Inhibitory
effect of bcl-2 on p53-mediated transactivation following
genotoxic stress, Oncogene 18 (1999) 297–304.
[41] S.A. Amundson, T.G. Meyers, A.J. Fornace, Roles for p53
in growth arrest and apoptosis: putting on the brakes after
genotoxic stress, Oncogene 17 (1998) 3287–3299.
[42] W.S. El-Diery, Regulation of p53 downstream genes, Semin.
Cancer Biol. 8 (1998) 345–357.
[43] H. Offer, R. Wolowicz, D. Matas, S. Blumenstein, Z.
Livneh, V. Rotter, Direct involvement of p53 in the base
excision repair pathway of the DNA repair machinery,
FEBS Lett. 450 (1999) 197–204.
[44] T. Grombacher, U. Eichorn, B. Kaina, p53 is involved in
regulation of the DNA repair gene O6-methylguanine–DNA
methyltransferase (MGMT) by DNA damaging agents,
Oncogene 17 (1998) 845–851.
[45] B.J. Hwang, J.M. Ford, P.C. Hanawalt, G. Chu, Expression
of the p48 xeroderma pigmentosum gene is p53-dependent
and is involved in global genomic repair, Proc. Natl. Acad.
Sci. 96 (1999) 424–428.
[46] J.M. Ford, P.C. Hanawalt, Li–Fraumeni syndrome fibroblasts
homozygous for p53 mutations are deficient in global DNA
repair but exhibit normal transcription-coupled repair and
enhanced UV resistance, Proc. Natl. Acad. Sci. 92 (1995)
8876–8880.
[47] J.M. Ford, P.C. Hanawalt, Expression of wild-type p53 is
required for efficient global genomic nucleotide excision
repair in UV-irradiated human fibroblasts, J. Biol. Chem.
272 (1997) 28073–28080.
[48] M.L. Smith, I.T. Chen, Q. Zhan, P.M. O’Conner, A.J.
Fornace, Involvement of the p53 tumor suppressor in repair
of UV-type DNA damage, Oncogene 10 (1995) 1053–1059.
[49] J.M. Ford, E.L. Baron, P.C. Hanawalt, Human fibroblasts
expressing the human papillomavirus E6 gene are deficient
in global genomic nucleotide excision repair and sensitive
to ultraviolet irradiation, Cancer Res. 58 (1998) 599–603.
[50] J.P. Therrien, R. Drouin, C. Baril, E.A. Drobetsky, Human
cells compromised for p53 function exhibit defective global
and transcription-coupled nucleotide excision repair, whereas
cells compromised for pRb function are defective only in
global repair, Proc. Natl. Acad. Sci. 96 (1999) 15038–15043.
[51] X. Wang, W.H. Yeh, L. Schaeffer, R. Roy, V. Moncollin,
J.M. Egly, Z. Wang, E.C. Friedberg, M.K. Evans, B.G. Taffe,
V.A. Bohr, G. Weeds, J.H.J. Hoeijmakers, K. Forrester,
C.C. Harris, p53 modulation of TFIIH-associated nucleotide
excision repair activity, Nat. Genet. 10 (1995) 188–195.
[52] G. Bakalkin, T. Yakovleva, G. Selivanova, K.P. Magnusson,
L. Szekely, E. Kiseleva, G. Klein, L. Terenius, K.G. Wiman,
![Page 15: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/15.jpg)
S.M. Morris /Mutation Research 511 (2002) 45–62 59
p53 binds single-stranded DNA ends and catalyzes DNA
renaturation and strand transfer, Proc. Natl. Acad. Sci. 91
(1994) 413–417.
[53] M. Reed, B. Woeler, P. Wang, Y. Wang, M.E. Anderson, P.
Tegtmeyer, The C-terminal domain of p53 recognizes DNA
damaged by ionizing radiation, Proc. Natl. Acad. Sci. 92
(1995) 9455–9459.
[54] L. Jayaraman, C. Prives, Activation of p53 sequence-specific
DNA binding by short single strands of DNA requires the
p53 C-terminus, Cell 81 (1995) 1021–1029.
[55] Y. Liu, M. Kulesz-Martin, p53 protein at the hub of cellular
DNA damage response pathways through sequence-specific
and non-sequence-specific DNA binding, Carcinogenesis
22 (2001) 851–860.
[56] Y. Saintigny, D. Rouilard, B. Chaput, T. Soussi, B.S. Lopez,
Mutant p53 proteins stimulate spontaneous and radiation-
induced intrachromosomal homologous recombination
independently of the alteration of the transactivation activity
and of the G1 checkpoint, Oncogene 18 (1999) 3553–3563.
[57] P. Bertrand, D. Rouillar, A. Boulet, C. Levalois, T. Soussi,
B.S. Lopez, Increase of spontaneous intrachromosomal
homologous recombination in mammalian cells expressing
a mutant p53 protein, Oncogene 14 (1997) 1117–1122.
[58] K.L. Meekel, W. Tang, L.A. Kachnic, C.M. Luo, J.S.
DeFrank, S.N. Powell, Inactivation of p53 results in high
rates of homologous recombination, Oncogene 14 (1997)
1847–1857.
[59] S. Buchop, M.K. Gibson, X.W. Wang, P. Wagner, H.W.
Sturzbecher, C.C. Harris, Interaction of p53 with the human
RAD51 protein, Nucl. Acids Res. 25 (1997) 3868–3874.
[60] C. Dudenhoffer, G. Rohaly, K. Will, W. Deppert, L.
Weismuller, Specific mismatch recognition in heteroduplex
intermediates suggests a role in fidelity control of homolo-
gous recombination, Mol. Cell. Biol. 18 (1998) 5332–5342.
[61] T. Mummenbrauer, F. Janus, B. Mueller, L. Weismulller, W.
Deppert, F. Grosse, p53 protein exhibits 3′–5′ exonuclease
activity, Cell 85 (1996) 1089–1099.
[62] F. Janus, N. Albrechtson, U. Knippschild, L. Wiesmuller,
F. Grosse, W. Deppert, Different regulation of the p53 core
domain activities 3′–5′ exonuclease and sequence-specific
DNA binding, Mol. Cell. Biol. 19 (1999) 2155–2168.
[63] Y. Saintigny, A. Dumay, S. Lambert, B.S. Lopez, A novel
role for the bcl-2 protein family: specific suppression of
the RAD51 recombination pathway, EMBO J. 20 (2001)
2596–2607.
[64] L.H. Thompson, D. Schild, The contribution of homologous
recombination in preserving genome integrity in mammalian
cells, Biochimie 81 (1999) 87–105.
[65] C. Gobert, A. Skladanowski, A.K. Larsen, The interaction
between p53 and DNA topoisomerase-I is regulated
differently in cells with wild-type and mutant p53, Proc.
Natl. Acad. Sci. 96 (1999) 10355–10360.
[66] A. Albor, S. Kaku, M. Kulesz-Martin, Wild-type and mutant
forms of p53 activate human topoisomersase I: a possible
mechanism for gain of function in mutants, Cancer Res. 58
(1998) 2091–2094.
[67] H.M. Smith, A.J. Grosovsky, Poly ADP-ribose-mediated
regulation of p53 complexed with topoisomerase I following
ionizing radiation, Carcinogenesis 29 (1999) 1439–1443.
[68] Y. Mao, S. Okada, L.S. Chang, M.T. Muller, p53 dependence
of topoisomerase I recruitment in vivo, Cancer Res. 60
(2000) 4538–4543.
[69] Q. Wang, G.P. Zambetti, D.P. Shuttle, Inhibition of DNA
topoisomerase IIa gene expression by the p53 tumor
suppressor, Mole. Cell. Biol. 17 (1997) 389–397.
[70] H. Yuwen, C.C. Hsia, Y. Nakashima, A. Evangelista, E.
Tabor, Binding of wild-type p53 by topoisomerase II and
overexpression of topoisomerase in human hepatocellular
carcinoma, Biochem. Biophys. Res. Commun. 234 (1997)
194–197.
[71] Y. Kwon, B.S. Shin, I.K. Chung, The p53 tumor suppressor
stimulates the catalytic activity of human topoisomerase IIa
by enhancing the rate of ATP hydrolysis, J. Biol. Chem.
275 (2000) 18503–18510.
[72] I.G. Cowell, A.L. Okorokov, S.A. Cutts, K. Padget, M.
Bell, J. Milner, C.A. Austin, Human topoisomerase IIa, and
IIb interact with the C-terminal region of p53, Exp. Cell
Res. 255 (2000) 86–94.
[73] P.N. Friedman, X. Chen, J. Bargonetti, C. Prives, The p53
protein is an unusually shaped tetramer that binds directly
to DNA, Proc. Natl. Acad. Sci. 90 (1993) 3319–3323.
[74] J. Milner, E.A. Medcalf, Cotranslation of activated mutant
p53 with wild type drives the wild-type p53 protein into
the mutant conformation, Cell 65 (1991) 765–774.
[75] J. Milner, E.A. Medcalf, A.C. Cook, Tumor suppressor
p53: analysis of wild-type and mutant p53 complexes, Mol.
Cell. Biol. 11 (1991) 12–19.
[76] A.C. Williams, J.C. Miller, T.J. Collard, T.S. Bracey, S.
Cosulich, C. Paraskeva, Mutant p53 is not fully dominant
over endogenous wild type p53 in a colorectal adenoma
cell line as demonstrated by induction of mdm2 protein
and retention of a p53 dependent G1 arrest after irradiation,
Oncogene 11 (1995) 141–149.
[77] G.P. Zambetti, A.J. Levine, A comparison of the biological
activities of wild-type and mutant p53, FASEB J. 7 (1993)
855–865.
[78] M. Levero, V. deLaurenzi, A. Costanzo, J. Gong, G. Melino,
J.Y.J. Wang, Structure, function and regulation of p63 and
p73, Cell Death Differ. 6 (1999) 1146–1153.
[79] C.A. Jost, M.C. Marin, W.G. Kaelin, p73 is a human
p53-related protein that can induce apoptosis, Nature 389
(1997) 191–194.
[80] M. Kaghad, H. Bonnet, A. Yang, L. Creacier, J.C. Biscan,
A. Valent, A. Minty, P. Chalon, J.M. Leilas, X. Dumont, P.
Ferrara, F. McKeon, D. Caput, Monoalleleically expressed
gene related to p53 at 1p36, a region frequently deleted
in neuroblastoma and other human cancers, Cell 90 (1997)
809–819.
[81] J. Zhu, J. Jiang, W. Zhous, X. Chen, The potential tumor
suppressor p73 differentially regulates cellular p53 target
genes, Cancer Res. 58 (1998) 5061–5065.
[82] R. Agami, G. Blandino, M. Oren, Y. Shaul, Interaction of
c-abl and p73a and their collaboration to induce apoptosis,
Nature 399 (1999) 809–813.
![Page 16: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/16.jpg)
60 S.M. Morris /Mutation Research 511 (2002) 45–62
[83] J. Gong, A. Constanzo, H.Q. Yang, G. Melinos, W.G.
Kaelin, M. Levero, J.J. Wang, The tyrosine kinase c-abl
regulates p73 in apoptotic response to cisplatin-induced
DNA damage, Nature 399 (1999) 806–809.
[84] A.M. Yuan, H. Shioya, T. Ishiko, X. Sun, J. Gu, Y. Huang,
H. Lu, S. Kharbanda, R. Weichselbaum, D. Kufe, p73 is
regulated by tyrosine kinase c-abl in the apoptotic response
to DNA damage, Nature 399 (1999) 814–817.
[85] T.S. Davison, C. Vagner, M. Kaghad, A. Ayed, D. Caput,
C.H. Arrowsmith, p73 and p63 are homotetramers capable
of weak heterotypic interactions with each other but not
with p53, J. Biol. Chem. 274 (1999) 18709–18714.
[86] C.J. DiComo, C. Gaiddon, C. Prives, p73 function is
inhibited by tumor-derived p53 mutants in mammalian
cells, Mol. Cell. Biol. 19 (1999) 1438–1449.
[87] M.C. Marin, C.A. Jost, L.A. Brooks, J. O’Nions, J.A. Tidy,
N. James, J.M. McGregor, C.A. Harwood, I.G. Yulug, K.H.
Vousden, M.J. Allday, B. Gusterson, S. Ikawa, P.W. Hinds,
T. Crook, W.G. Kaelin, A common polymorphism acts as
an intragenic modifier of mutant p53 behavior, Nat. Genet.
25 (2000) 47–54.
[88] A. Yang, M. Kaghad, Y. Wang, E. Gillet, M.D. Fleming,
V. Dotsch, N.C. Andrews, D. Caput, F. McKeon, p63,
a p53 homologue at 3q27-29, encodes multiple products
with transactivating, death-inducing, and dominant-negative
activities, Mol. Cell 2 (1998) 305–316.
[89] C.J. Sherr, J.D. Weber, The ARF/p53 pathway, Curr. Opin.
Genet. Dev. 10 (2000) 94–99.
[90] M. Ljungman, Dial 9-1-1 for p53: mechanisms of p53
activation by cellular stress, Neoplasia 2 (2000) 208–225.
[91] M. Ashcroft, K.H. Vousden, Regulation of p53 stability,
Oncogene 18 (1999) 7637–7643.
[92] N.E. Sharpless, R.A. Pinho, The INK4A/ARF locus and its
two gene products, Curr. Opin. Genet. Dev. 9 (1999) 22–30.
[93] W. Tao, A.J. Levine, p19ARF stabilizes p53 by blocking
nucleo-cytoplasmic shuttling of mdm2, Proc. Natl. Acad.
Sci. 96 (1999) 6937–6941.
[94] J.D. Weber, M.L. Kuo, B. Bothner, E.L. Di Giammarino,
R.W. Kriwacki, M.F. Roussel, C.J. Sherr, Cooperative
signals governing the ARF–mdm2 interaction and nuclear
localization of the complex, Mol. Cell. Biol. 20 (2000)
2517–2528.
[95] C.J. Sherr, Tumor surveillance via the ARF-p53 pathway,
Genes Dev. 12 (1998) 2984–2991.
[96] D.P. Lane, L.V. Crawford, T-antigen is bound to host protein
in SV40-transformed cells, Nature 278 (1979) 261–263.
[97] F. Mantovani, L. Banks, The interaction between p53 and
papillomaviruses, Semin. Cancer Biol. 9 (1999) 387–395.
[98] H. Zimmerman, R. Degenkolbe, H.U. Bernard, M.J.
O’Conner, The human papillomavirus Type 16 E6
oncoprotein can down-regulate p53 activity by targeting the
transcriptional coactivator CBP/p300, 1999.
[99] M. Thomas, D. Pim, L. Banks, The role of E6–p53 inter-
action in the molecular pathogenesis of HPV, Oncogene 18
(1999) 7690–7700.
[100] M.E.D. Martin, A.J. Berk, Adenovirus E1B 55K represses
p53 activation in vitro, J. Virol. 72 (1998) 3146–3154.
[101] P.H. Gallimore, P.S. Lecane, S. Roberts, S.M. Rookes,
R.J.A. Grand, J. Parkhill, Adenovirus type 12 early region
1B 54K protein significantly extends the life span of normal
mammalian cells in culture, J. Virol. 71 (1997) 6629–6640.
[102] S.A. Amundson, F. Xia, K. Wolfson, H.L. Liber, Different
cytotoxic and mutagenic responses induced by X-rays in
two human lymphoblastoid cell lines derived from a single
donor, Mutat. Res. 286 (1993) 233–241.
[103] A.J. Grosovsky, B.N. Walter, C.R. Giver, DNA-sequence
specificity of mutations at the thymidine kinase locus,
Mutat. Res. 289 (1993) 231–243.
[104] F. Xia, S.A. Amundson, J.A. Nickloff, H.L. Liber, Different
capacities for recombination in closely related human
lymphoblastoid cell lines with different mutational responses
to X-irradiation, Mol. Cell. Biol. 14 (1994) 5850–5857.
[105] W. Zhen, C.M. Denault, K. Loviscek, S. Walter, L. Geng,
A.T.M. Vaughn, The relative radiosensitivity of TK6 and
WIL2-NS lymphoblastoid cells derived from a common
source is primarily determined by their p53 mutational
status, Mutat. Res. 346 (1995) 85–92.
[106] F. Xia, X. Wang, Y.H. Wang, N.M. Tsang, D.W. Yandell,
K.T. Kelsey, H.L. Liber, Altered p53 status correlated with
differences in sensitivity to radiation-induced mutation and
apoptosis in two closely related human lymphoblast lines,
Cancer Res. 55 (1995) 12–15.
[107] M. Honma, M. Hayashi, T. Sofuni, Cytotoxic and mutagenic
responses to X-rays and chemical mutagens in normal and
p53-mutated human lymphoblastoid cells, Mutat. Res. 374
(1995) 89–98.
[108] M. Honma, L.-S. Zhang, M. Hayashi, K. Takeshita, Y.
Nakagawa, N. Tanaka, T. Sofuni, Illegitimate recombination
leading to allelic loss and unbalanced translocation in
p53-mutated human lymphoblastoid cells, Mol. Cell. Biol.
17 (1997) 4774–4781.
[109] F. Xia, H.L. Liber, The tumor suppressor p53 modifies
mutational processes in a human lymphoblasotid cell line,
Mutat. Res. 373 (1997) 87–97.
[110] M. Honma, M. Momose, H. Tanabe, H. Sakamoto, Y.
Yu, J.B. Little, T. Sofuni, M. Hayashi, Requirement of
wild-type p53 protein for maintenance of chromosomal
integrity, Mol. Carcinog. 28 (2000) 203–214.
[111] Y. Yu, C.Y. Li, J.B. Little, Abrogation of p53 function by
HPV16E6 gene delays apoptosis and enhances mutagenesis
but does not alter radiosensitivity in TK6 human
lymphpblastoid cells, Oncogene 14 (1997) 1661–1667.
[112] Y.Y.E. Chuang, Q. Chen, H.L. Liber, Radiation-induced
mutations at the autosomal thymidine kinase locus are not
elevated in p53-null cells, Cancer Res. 59 (1999) 3073–3076.
[113] C.L. Crespi, W.G. Thilly, Assay for gene mutation in a
human lymphoblast line, AHH-1, competent for xenobiotic
metabolism, Mutat. Res. 128 (1984) 221–230.
[114] K.L. Dobo, C.R. Giver, D.A. Eastmond, H.S. Rumbos, A.J.
Grosovsky, Extensive loss of heterozygosity accounts for
differential mutation rate on chromosome 17q in human
lymphoblasts, Mutagenesis 10 (1995) 53–58.
[115] S.M. Morris, M.G. Manjanatha, S.D. Shelton, O.E. Domon,
L.J. McGarrity, D.A. Casciano, A mutation in the p53 tumor
![Page 17: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/17.jpg)
S.M. Morris /Mutation Research 511 (2002) 45–62 61
suppressor gene of AHH-1 TK+/− human lymphoblastoid
cells, Mutat. Res. 356 (1996) 129–134.
[116] S.M. Morris, J.J. Chen, O.E. Domon, L.J. McGarrity, M.E.
Bishop, M.G. Manjanatha, D.A. Casciano, p53, mutations
and apoptosis in genistein-exposed lymphoblastoid cells,
Mutat. Res. 405 (1998) 41–56.
[117] O.E. Domon, L.J. McGarrity, M. Bishop, M. Yoshioka,
J.J. Chen, S.M. Morris, Evaluation of the genotoxicity of
the phytoestrogen, coumestral, in AHH-1 TK+/− human
lymphoblasotid cells, Mutat. Res. 474 (2001) 129–137.
[118] K.L. Dobo, D.A. Eastmond, A.J. Grosovsky, The influence
of cellular apoptotic capacity on N-nitrosodimethylamine-
induced loss of heterozygosity mutations in human cells,
Carcinogenesis 18 (1997) 1701–1707.
[119] H. Masuda, C. Miller, H.P. Koefller, H. Battifor, M.J.
Cline, Rearrangement of the p53 gene in human osteogenic
sarcomas, Proc. Natl. Acad. Sci. 84 (1987) 7716–7719.
[120] N. Yamagishi, J. Miyakoshi, H. Takebe, Decrease in the
frequency of X-ray-induced mutation by wild-type p53
protein in human osteosarcoma cells, Carcinogenesis 18
(1997) 695–700.
[121] T. Yagi, K. Mohri-Nakanishi, T. Matsuda, N. Yamagishi, J.
Miyakoshi, H. Takebe, Reduced UV-induced mutations in
human osteosarcoma cells stability expressing transfected
wild-type p53 cDNA, Cancer Lett. 123 (1998) 71–76.
[122] P.A. Havre, J. Yuan, L. Hedrick, K.R. Cho, P.M. Glazer, p53
inactivation by HPV16E6 results in increased mutagenesis
in human cells, Cancer Res. 55 (1995) 4420–4424.
[123] J. Yuan, T.M. Yeasky, P.A. Havre, P.M. Glazer, Induction of
p53 in mouse cells decreases mutagenesis by UV radiation,
Carcinogenesis 16 (1995) 2295–2300.
[124] E.N. Phillips, D. Gebow, H.L. Liber, Spectra of X-ray-
induced and spontaneous intragenic HPRT mutations in
closely related human cells differentially expressing the
p53 tumor suppressor gene, Radiat. Res. 147 (1997)
138–147.
[125] L.A. Donehower, M. Harvey, B.L. Slagle, M.J. McArthur,
C.A. Montogomery, J.S. Butel, A. Bradley, Mice deficient
for p53 are developmentally normal but susceptible to
spontaneous tumors, Nature 356 (1992) 215–221.
[126] T. Jacks, L. Remington, B.O. Williams, E.M. Schmitt, S.
Halachmi, R.T. Bronson, R.A. Weinberg, Tumor spectrum
analysis in p53-mutant mice, Curr. Biol. 4 (1994) 1–7.
[127] C.A. Purdie, D.J. Harrison, A. Peter, L. Dobbie, S. White,
S.E.M. Howie, D.M. Salter, C.C. Bird, A.H. Wyllie, M.L.
Hooper, A.R. Clarke, Tumor-incidence, spectrum and ploidy
in mice with a large deletion in the p53 gene, Oncogene 9
(1994) 603–609.
[128] K. MacLeod, T. Jacks, Insights into cancer from transgenic
mouse models, J. Pathol. 187 (1999) 43–60.
[129] S. Venkatachalam, Y.P. Shi, S.N. Jones, H. Vogel, A.
Bradley, D. Pinkel, L.A. Donehower, Retention of wild-type
p53 in tumors from p53 heterozygous mice: reduction of
p53 dosage can promote cancer formation, EMBO J. 17
(1998) 4657–4667.
[130] H. Nishino, A. Knoll, B. Buettner, C.S. Fisk, Y. Maruta,
J. Haavik, S.S. Somer, p53 wild-type and p53 nullizygous
Big Blue© transgenic mice have similar frequencies and
patterns of observed mutation in liver, spleen and brain,
Oncogene 11 (1995) 263–270.
[131] V.L. Buettner, H. Nishino, J. Haavik, A. Knoll, K. Hill,
S.S. Somer, Spontaneous mutation frequencies and spectra
in p53+/+ and p53−/− mice: a test of the guardian
of the genome hypothesis in the Big Blue© transgenic
mouse mutation detection systems, Mutat. Res. 379 (1997)
13–20.
[132] V.L. Buettner, K.A. Hill, H. Nishino, D.J. Schaid, C.S.
Frisk, S.S. Somer, Increased mutation frequency and altered
spectrum in one of four lymphomas derived from tumor
prone p53/Big Blue© double transgenic mice, Oncogene 13
(1996) 2407–2413.
[133] A.T. Sands, M.B. Suragkar, A. Sanchez, J.E. Marth, L.A.
Donehower, A. Bradley, p53 deficiency does not affect the
accumulation of point mutations in a transgene target, Proc.
Natl. Acad. Sci. 92 (1995) 8517–8521.
[134] P.K. Gupta, A. Sahota, S. Bye, S. Boyadjiev, C. Shao,
J.P. O’Neill, R.J. Albertini, P.J. Stambrook, J.A. Tischfield,
High frequency in vivo loss of heterozygosity is primarily
a consequence of mitotic recombination, Cancer Res. 57
(1997) 1188–1193.
[135] C. Shao, L. Deng, O. Henagariu, L. Liang, N. Raikwar, A.
Sahota, P.J. Stambrook, J.A. Tischfield, Mitotic recombina-
tion produces the majority of recessive fiborblast variants
in heterozygous mice, Proc. Natl. Acad. Sci. 96 (1999)
9230–9235.
[136] C. Shao, L. Deng, O. Henegariu, P.J. Stambrook, J.A.
Tischfield, Chromosome instability contributes to loss of
heterozygosity in mice lacking p53, Proc. Natl. Acad. Sci.
97 (2000) 7405–7410.
[137] M.H. Brilliant, Y. Gondo, E.M. Eicher, Direct molecular
identification of the mouse pink-eyed unstable mutation by
genome scanning, Science 266 (1991) 1572–1576.
[138] Y. Gondo, J.M. Gardner, Y. Nakatsu, D. Durham-Pierre,
S.A. Deaveau, C. Kuper, M.H. Brilliant, High frequency
genetic reversion mediated by a DNA duplication: the
mouse pink-eyed unstable mutation, Proc. Natl. Acad. Sci.
90 (1993) 297–301.
[139] R.H. Schiestl, F. Koghali, N. Carls, Reversion of the mouse
pink-eyed unstable mutation induced by low doses of
X-rays, Science 266 (1994) 1573–1576.
[140] J. Aubrecht, M.B. Secretan, A.J.R. Bishop, R.H. Schiestl,
Involvement of p53 in X-ray induced intrachromosomal
recombination in mice, Carcinogenesis 20 (1999) 2229–
2236.
[141] A. De Vries, C.T.M. van Oostrom, F.M.A. Hofhuis, P.M.
Dortant, R.J.W. Berg, F.R. DeGruiji, P.W. Wester, C.F. Van
Kreijl, P.J.A. Capel, H. van Steeg, S.J. Verbeel, Increased
susceptibility to ultraviolet-B and carcinogens of mice
lacking the DNA excision repair gene Xpa, Nature 377
(1995) 169–173.
[142] C.T.M. van Oostrom, M. Boeve, J. van den Berg, A.
De Vries, M.E.T. Dolle, R.B. Beems, C.F. Van Kreijl,
J. Vijg, H. van Steeg, Effect of heterozygous loss of
p53 on benzo(a)pyrene induced mutations and tumors in
![Page 18: A Role for p53 in the Frequency and Mechanism of Mutation, 2002](https://reader035.vdocuments.site/reader035/viewer/2022073116/552cdf4b4a79597f578b457f/html5/thumbnails/18.jpg)
62 S.M. Morris /Mutation Research 511 (2002) 45–62
DNA-repair-deficient Xpa mice, Environ. Mol. Mutagen.
34 (1999) 124–130.
[143] H. van Steeg, The role of excision repair and loss of p53 in
mutagenesis and carcinogenesis, Toxicol. Lett. 120 (2001)
209–219.
[144] S.D. Griffiths, A.R. Clarke, L.E. Healy, G. Ross, A.M. Ford,
M.L. Hooper, A.H. Wyllie, M. Greaves, Absence of p53
permits propagation of mutant cells following genotoxic
damage, Oncogene 14 (1997) 523–531.
[145] P. Li, P.D. Sutphin, D. Schwartz, D. Matas, N. Almog,
R. Wolkowicz, N. Goldfinger, H. Pei, M. Prokocimer,
V. Rotter, Mutant p53 protein expression interferes with
p53-independent apoptotic pathways, Oncogene 16 (1998)
3269–3277.
[146] C. Cherbonnel-Lassere, S. Gauny, A. Kroneneberg, Suppres-
sion of apoptosis by bcl-2 and bcl-xl promotes susceptibility
to mutagenesis, Oncogene 19 (1996) 1489–1497.
[147] C. Cherbonnel-Lassere, M.K. Dosanjh, Suppression of
apoptosis by overexpression of bcl-2 or bcl-xl promotes
survival and mutagenesis after oxidative damage, Biochimie
79 (1997) 613–617.
[148] V.N. Doborovolsky, D.A. Casciano, R.H. Heflich, TK+/−
mouse model for detecting in vivo mutation in an endoge-
nous, autosomal gene, Mutat. Res. 423 (1999) 125–136.
[149] G. Liu, T.J. McDonnell, R.M. de Oca Luna, M. Kapoor,
B. Mims, A.K. El-Naggar, G. Lozano, High metastatic
potential in mice inheriting a targeted p53 missense muta-
tion, Proc. Natl. Acad. Sci. 97 (2000) 4174–4179.