Occurrence of H-ras codon 61 CAA to AAA mutation during mouse liver tumor progression

Barbara L. Parsons1,3, Sandra J. Culp2, Mugimane G. Manjanatha1 and Robert H. Heflich1

1 Division of Genetic and Reproductive Toxicology and
2 Division of Biochemical Toxicology, HFT-120, National Center for Toxicological Research, 3900 NCTR Road, Jefferson, AR 72079, USA


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The initiating mutations of a tumor are present in each of the cancerous cells comprising the tumor. Identification and measurement of the subsequent mutations that occur during tumor progression, however, requires mutation detection in a smaller subset of the tumor cells. In this study, allele-specific competitive blocker PCR (ACB-PCR), a genotypic selection method with the sensitivity to detect a specific point mutation in the presence of a 105-fold excess of wild-type DNA sequence, was used to measure H-ras codon 61 CAA to AAA mutation in mouse liver tumors that did not have this mutation as an initiating event. Twenty-one spontaneous or chemically induced mouse liver tumors, negative for the H-ras codon 61 CAA to AAA mutation by DNA sequencing or denaturing gradient gel electrophoresis, were analyzed for this mutation by ACB-PCR. The mutation was detected at some level in 71% of these tumors. The mutation was detected in adenomas and carcinomas more frequently (13 of 14 tumors) and at significantly higher mutant fractions than it was detected in histiocytic sarcomas (1 of 5 tumors). These data indicate that the same oncogenic point mutation that can be identified as a tumor-initiating event based on its clonal amplification in a tumor can also be present in only a small sub-population of tumor cells where the mutation must have been fixed at a later stage in tumor development. The occurrence of a mutation as a primary or secondary event probably reflects the stochastic nature of mutation and is likely to be affected by the mutation rate for each target site.

Abbreviations: ACB-PCR, Allele specific competitive blocker PCR; bp, basepair, DGGE, denaturing gradient gel electrophoresis; PCR, polymerase chain reaction.


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Models of multistage carcinogenesis have been developed for a number of different tumor types (13). According to these models, cells acquire multiple mutations during tumor progression from pre-malignant lesions to small adenomas and eventually to metastatic carcinomas (4). Point mutations in oncogenes are often early events in this process, followed by loss of tumor suppressor genes, microsatellite instability, loss of heterozygosity, and aneuploidy. The additional genetic changes acquired by initiated cells result in the new cellular phenotypes and capabilities that lead to malignancy (5). Thus, mutations are regarded as causative events in carcinogenesis.

Predictions have been made regarding the number of genetic changes required to produce a malignant phenotype. These predictions, which are primarily based on age-dependent tumor incidences and vary in terms of their underlying assumptions and the type of tumor being modeled, suggest that between two and seven mutations are necessary (6,7). By sequencing tumor DNA, many `tumor-initiating' mutations have been discovered. Nearly 100 different oncogenes and >20 tumor suppressor genes have now been identified and many of these loci may be affected by a number of different activating mutations (4). While it is clear that there are many different mutational targets, it is also known that mutation at any given site is a rare event. Thus, the acquisition of genetic change during tumorigenesis is a stochastic process and many different combinations of these rare genetic events are possible. A consequence of this stochastic process is that tumors are genetically and phenotypically heterogeneous.

Although rodent cells are more susceptible to transformation than human cells, the concept of multistage carcinogenesis applies equally well to humans and rodents (8). Consequently, the mouse liver tumor model can be used to study both multistage carcinogenesis and tumor progression (9). One distinguishing feature of the mouse liver tumor model is the predominance of a few specific point mutations. Mutations at H-ras codon 61 are the most common mutations observed in mouse liver tumors; they appear in ~56% of spontaneous B6C3F1 liver tumors (10). This high frequency of H-ras point mutation is mouse strain-specific, the result of selective breeding for liver tumor-sensitive mouse strains. The H-ras codon 61 CAA to AAA mutation accounts for ~60% of these codon 61 mutations. Thus, it can be calculated that this one specific point mutation appears in ~34% of all spontaneous B6C3F1 mouse liver tumors. Furthermore, treatment with a number of different chemicals raises the frequency of this point mutation even higher (10). The fact that this point mutation occurs so frequently as a primary initiating mutation raises the possibility that this mutation might also occur relatively frequently as a subsequent mutation. If so, then the H-ras codon 61 CAA to AAA mutation might be a useful biomarker with which to study progression of mouse liver tumors.

Because initiating mutations are the mutations that lead to a transformed phenotype and uncontrolled cell proliferation, they are expected to be present in all the transformed cells of a particular tumor. Although excised tumors often contain some normal cells, the fraction of cells containing the initiating mutation should approach unity. In contrast, mutations that arise after the initiating events, hereafter referred to as `secondary mutations,' should be present in a smaller fraction of the transformed cells of a tumor (11). Thus, these secondary mutations that occur during tumor progression can be distinguished based on their prevalence in tumor DNA. In fact, it can be assumed that the smaller the mutant fraction of a particular tumor-associated mutation the later in tumor progression that mutation occurred.

Once a secondary mutation has occurred, it can be amplified clonally in the tumor. Clonal amplification and the use of a frequently mutated target are factors that will facilitate the detection of the secondary mutational events that occur during tumor progression. On the other hand, the presence of non-transformed cell types in excised tumor tissue and the fact that many different secondary events could lead to tumor outgrowth are factors that make the measurement of such secondary mutations difficult. Clearly, specialized techniques are needed to detect these rare and possibly low level mutational events. Genotypic selection, the DNA-based detection of rare mutation, is an ideal approach for this application (12). Allele-specific competitive blocker-PCR (ACB-PCR) is a type of genotypic selection that relies on the selective PCR amplification of mutant alleles (13). The ACB-PCR method was adapted to detect the mouse H-ras codon 61 CAA to AAA mutation and modified such that mutant fractions as low as 10–5 could be distinguished from wild-type samples (14). In this study, ACB-PCR was used to determine whether mouse liver tumors that did not have the H-ras codon 61 CAA to AAA mutation as an initiating event acquired this mutation during liver tumor progression.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
DNA standards. H-ras codon 61 mutant (AAA) and wild-type (CAA) plasmids (encompassing mouse H-ras exon 1, intron 1, and part of exon 2) were constructed as previously described (15). A Pvu II–Hind III restriction fragment was gel-purified from each plasmid (Geneclean Spin Kit, Bio 101, Vista, CA) and quantified by SYBR Green I staining (Molecular Probes, Eugene, OR) relative to a Low DNA Mass® Ladder (Gibco BRL, Rockville, MD) using a Molecular Dynamics FluorImager (Amersham Pharmacia Biotech, Piscataway, NJ). The purified restriction fragments (3.75 ng) were used as templates for polymerase chain reaction (PCR) to amplify 189-basepair (bp) fragments corresponding to either the mutant or wild-type H-ras DNA sequence. Each 100 µl PCR reaction contained: 10 mM KCl, 10 mM (NH4)2SO4, 20 mM Tris–HCl (pH 8.75), 2 mM MgSO4, 0.1% Triton X-100, 100 µg/ml bovine serum albumin, 750 µM dNTPs, 1.5 µM primer TR3 (5'-TTCTGTGGATTCTCTGGT-3'), 1.5 µM primer H2 (5'-GTGCG-CATGTACTGGTCCCG-3'), and 3.75 units of Pfu DNA polymerase (Stratagene, La Jolla, CA). The PCR products were isolated from agarose gels using a Geneclean Spin Kit and frozen (in single-use aliquots) as DNA copy number standards. The DNA concentration of these aliquots was repeatedly determined relative to the Low DNA Mass Ladder until three different determinations that varied by <=10% were obtained. Cycling conditions were 1 min at 94°C, 5 min at 48°C, and 1 min at 72°C for one cycle followed by 28 cycles of 1 min at 94°C, 1 min at 48°C, and 2 min at 72°C.

Tumor DNA isolation and PCR amplification of tumor H-ras DNA. Mouse liver tumor DNAs were collected that were negative for the H-ras codon 61 mutation as determined by DNA sequencing (16) or denaturing gradient gel electrophoresis (DGGE) (17). DNA was isolated from one-half of these tumors as previously described (tumors M4, M5, M6, 464, 466, 467, 494 (16)) (all other tumors (17)); the other half being used for histological examination. The same 189 bp region encompassing the H-ras intron 1-exon 2 junction was PCR amplified from these tumor DNA samples. 1.5 µg of each tumor DNA was used as template (this corresponds to ~4.5 x 105 copies of H-ras DNA sequence (12)). Except for the template DNA, PCR conditions were identical to those described above for the preparation of H-ras mutant and wild-type copy number standards.

Allele-specific competitive blocker-PCR (ACB-PCR). Varying mixtures of mutant and wild-type copy number standard DNAs were prepared. Each DNA mixture contained 90 pg of wild-type sequence (4.6 x 108 copies). Different amounts of diluted mutant DNA were added to give mutant fractions ranging from 10-1 to 10-5. A control sample comprised of 90 pg of wild-type sequence without any added mutant sequence was always included. The ACB-PCR reactions contained three different primers (Sigma Genosys, The Woodlands, TX); the mutant specific primer (TR10, 5'-ATGGCACTATACTCTTGTCT-3'), the upstream primer (TR 31, 5'-TGGGGAGACATGTCTACTG-3'), and a blocker primer (TR27, 5'-ATGGCACTATACTCTTCTAG-3'). The blocker primer carries a 3'-terminal dideoxy guanosine residue and, therefore, cannot be extended. The mutant-specific and blocker primers were gel-purified. The DNA mixtures were analyzed in 50 µl ACB-PCR reaction mixtures containing: 10 mM KCl, 10 mM Tris–HCl (pH 8.3), 1.5 mM MgCl2, 1 mg/ml Triton X-100, 0.1 mg/ml gelatin, 20 µM of each dNTP, 400 nM TR10, 330 nM TR31, 200 nM TR27, 0.06 units of Perfect Match PCR Enhancer (Stratagene), and three units of Taq DNA polymerase Stoffel Fragment (PE Applied Biosystems, Foster City, CA). An enzyme mixture was prepared by diluting the Taq DNA polymerase Stoffel fragment and Perfect Match PCR Enhancer into the reaction mixture. DNA-containing reaction mixtures (45 µl) were preheated at 94°C for 2 min. The enzyme mixture was heated at 94°C for the last 20 s of that incubation and then 5 µl of the enzyme mixture were added to each reaction. The cycling conditions were 30 s at 94°C, 45 s at 46°C, and 1 min at 72°C, for 35 cycles.

Analysis of ACB-PCR products. Equal volumes of ACB-PCR products were analyzed on 8% polyacrylamide gels along with a 50 bp DNA Ladder (Gibco-BRL). The gels were stained with SYBR Green I. DNAs in the gels were detected using a FluorImager and the amount of the 60 bp, H-ras-specific PCR product was quantified using ImageQuaNT software (Molecular Dynamics, Amersham Pharmacia Biotech). The fluorescence determinations from the mutant fraction standards were used to construct a standard curve from which the H-ras codon 61 CAA to AAA mutant fractions in the tumor DNA samples were determined.


    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Livers tumors that had been characterized as H-ras negative were used in this investigation. These liver tumors were derived from two separate studies. In one study (Study #1), newborn male B6C3F1 and CD-1 mice were treated with 6-nitrochrysene, 4-aminobiphenyl, or the dimethylsulfoxide (DMSO) vehicle (16). The tumors were harvested when the animals were 12 months of age. DNAs from these liver tumors were used as templates to PCR amplify H-ras gene sequences. The PCR products were analyzed by fluorescent cycle-sequencing to determine which tumors carried H-ras mutation. Only liver tumor DNAs that were negative for H-ras mutation as determined by DNA sequencing were included in the present study. All of the tumors obtained from this study had been subject to pathological examination and identified as hepatocellular adenomas. The specific treatments and doses that induced each adenoma from Study #1 are given in Table IGo.


View this table:
[in this window]
[in a new window]
 
Table I. ACB-PCR analysis of H-ras Codon 61 CAA to AAA mutant fraction in seven mouse liver tumors induced as part of Study #1
 
In a second study (Study #2), tumors were harvested at the end of a 2-year bioassay in which 5- to 6-week-old female B6C3F1 mice were placed on a diet containing one of two different coal tar mixtures, benzo[a]pyrene, or feed treated with the acetone vehicle (18). Again, tumor DNAs were used as template to produce H-ras-specific PCR products which then were screened for mutation by DGGE (17). Liver tumor DNAs from Study #2 that were negative for H-ras mutation as determined by DGGE were included in the present study. The specific treatments that induced each tumor from Study #2 are given in Table IIGo, along with the tumor type determined by pathological examination.


View this table:
[in this window]
[in a new window]
 
Table II. ACB-PCR analysis of H-ras Codon 61 CAA to AAA mutant fraction in 14 liver tumors from female B6C3F1 mice induced as part of Study #2
 
Because of its high fidelity, Pfu DNA polymerase was used in the present study to generate first round PCR products from the 21 different tumor DNA samples. Then, ACB-PCR was performed as described in Materials and methods to specifically amplify mouse H-ras codon 61 CAA to AAA mutant DNA sequence from the 21 different first round PCR products. This mutant-specific amplification was achieved through the use of three different PCR primers and conditions that magnify the effects of 3'-terminal mismatches. The design of the PCR primers that leads to this selective PCR amplification is described in Figure 1Go. Because the mutant-specific primer has more 3'-terminal mismatches relative to the wild-type sequence than the mutant sequence, it preferentially amplifies mutant DNA sequence. The blocker primer has more 3'-terminal mismatches relative to the mutant sequence than the wild-type sequence, so it preferentially anneals to the wild-type DNA sequence but produces no product because it carries a 3'-terminal, chain terminator.



View larger version (34K):
[in this window]
[in a new window]
 
Fig. 1. PCR priming strategy for ACB-PCR. The basis for the specific amplification of mutant allele in ACB-PCR is the differential annealing of the mutant-specific and blocker primers to the mutant and wild-type templates, with extension of only the mutant-specific primer. Each of the four possible template/primer pairs is depicted (template above, primer below). The mutant-specific primer (MSP) and the blocker primer (BP) each have a mismatch in the 3'-penultimate position relative to either the mutant or wild-type template (mismatched bases are indicated in bold). The 3'-terminal base of the mutant specific primer is a T, which is complementary to the mutant A. The 3'-terminal base of the blocker primer is a dideoxy G. It is complementary to the wild-type base (C) but cannot be extended.

 
Three ACB-PCR experiments were performed in order to analyze the 21 different tumor DNA samples (Figure 2Go). In each experiment, PCR products synthesized from the tumor DNAs were analyzed in parallel with the mutant fraction standards and a no-mutant control. The mutant fraction standards were used to construct a standard curve relating fluorescence to H-ras mutant fraction. Using the standard curve, the tumor H-ras mutant fractions were calculated. The level of H-ras codon 61 CAA to AAA mutation detected in each tumor from Study #1 is given in Table IGo and the levels of mutation detected in tumors from Study #2 are given in Table IIGo. The mutation was detected in 15 of the 21 liver tumor DNAs analyzed (71%). Because the mutant fraction was quantified, the liver tumors can be stratified in terms of their H-ras codon 61 CAA to AAA mutant fractions. Two tumors had mutant fractions between 10-1 and 10-2, one tumor had a mutant fraction between 10-2 and 10-3, seven tumors had mutant fractions between 10-3 and 10-4, and five had mutant fractions between 10-4 and 10-5. When the detection of the H-ras mutation was examined in relation to tumor type, some differences were noted. An H-ras mutant fraction was detected in 11 of the 12 adenomas and both of the carcinomas included in this study, but was detected in only one of the five histiocytic sarcomas analyzed (Table IIIGo). In addition, when the mutant fractions detected in adenomas and carcinomas from Study #2 are compared with those of the histiocytic sarcomas from the same study, the adenomas/carcinomas showed significantly higher mutant fractions (t-test, P = 0.028, assigning mutant fractions of 9.9 x 10-6 to H-ras-negative samples).



View larger version (96K):
[in this window]
[in a new window]
 
Fig. 2. ACB-PCR detection of the mouse H-ras codon 61 CAA to AAA mutation. Twenty-one different mouse liver tumor DNAs, which were negative for the H-ras codon 61 CAA to AAA mutation by either DNA sequencing or DGGE, were analyzed by ACB-PCR. Each panel shows a subset of the PCR products amplified from the tumor DNA samples that were analyzed by ACB-PCR in parallel with mutant fraction standards and a no-mutant control. From left to right, the samples shown in (A) correspond to tumors 494, M4, M5, M6, 464, and 467; the samples shown in (B) correspond to tumors 466, L685, L727, L472, L554, L589, L678, and 170; the samples in (C) correspond to tumors 314, 343, 208, 400, 425, 393, and 789. The mutant fraction was calculated based on the fluorescence of the 60 bp H-ras specific PCR product relative to that of the mutant fraction standards.

 

View this table:
[in this window]
[in a new window]
 
Table III. Frequency of mutant H-ras detection in relation to tumor type.
 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The detection of activating ras gene mutations in pre-neoplastic and neoplastic foci and their detection in relatively large proportions of tumors from a number of different tissues has demonstrated that ras point mutation is frequently an early event in carcinogenesis (19,20). The role of ras point mutation in the later stages of tumor progression, however, is not understood, probably due to the lack of experimental approaches to address the question. This paper describes a novel approach for studying the accumulation of an oncogenic point mutation during mouse liver tumor progression. The approach was based on first identifying tumors that do not carry the H-ras codon 61 CAA to AAA mutation as an initiating lesion and then using a sensitive genotypic selection method (ACB-PCR) to detect the mutation as a sub-population of the tumor DNA.

In Study #1, the determination that the tumors did not carry an initiating H-ras mutation was based on DNA sequencing. The limit of sensitivity for detecting this mutation by DNA sequencing is ~1.25 x 10-1 (as the mutation is barely detectable in a mixture composed of one mutant allele per seven wild-type alleles, data not shown). All the H-ras mutant fractions detected by ACB-PCR were clearly below the limit of detection by DNA sequencing. Tumors derived from Study #2 were characterized by DGGE (17). DGGE is a more sensitive mutation detection method than DNA sequencing. Reports of DGGE sensitivity range between 10-2 and 10-3 (12). Again, all the tumors from Study #2 analyzed by ACB-PCR had mutant fractions below the sensitivity of the screening assay. Tumors generated in Study #2 that had non-initiating, but relatively high (>10-3) H-ras codon 61 AAA mutant fractions would have been identified by DGGE and, consequently, excluded from the present study. This may explain, at least in part, why none of the tumors characterized by DGGE had H-ras codon 61 mutant fractions >1.5 x 10-4 (Table IIGo).

The ACB-PCR technique detected H-ras mutation in 15 of the 21 tumors analyzed, however, before it can be concluded that these were secondary mutations in clonally-expanding tumor cells, several other possibilities must be excluded. First, it must be known that the mutations detected by ACB-PCR were not initiating mutations. Secondly, the possibility that the mutant fractions were assay-generated or represent secondary mutational events in contaminating normal tissue must be considered.

Did ACB-PCR simply detect initiating mutations? The mutant fraction measurements are sufficiently low that the H-ras mutations measured could not represent the tumor-initiating mutations expected to be present in every cell of the tumor. A possibility that was considered is that some tumor initiating mutations were not identified because they were contaminated with normal tissue, thereby lowering the apparent mutant fraction. This possibility was discounted, however, because even the tumors with the highest mutant fractions (~5 x 10-2, Table IGo) still had a 20-fold excess of wild-type relative to mutant sequence.

Once it is concluded that the tumors did not contain the H-ras mutation as an initiating mutation, the next concern is whether the mutations detected were assay-generated. The idea that the mutations detected by ACB-PCR can be attributed to PCR-associated, DNA polymerase-induced errors also can be excluded. The rate at which Pfu DNA polymerase makes this particular basepair substitution error is 8 ± 3 x 10-7 per basepair duplication, which translates into post-PCR mutant fractions of ~10-6 (21). The mutant fractions that were measured were all 10-5 or higher, as ACB-PCR is not sensitive enough to detect mutations below 10-5. Furthermore, any Pfu-induced errors that were generated would become part of the assay's background signal because the mutant fraction standards were synthesized using the same PCR conditions as those used to generate the first round PCR products from the tumor DNAs.

A further concern is whether the mutations detected were secondary mutations in the excised tissue but were not part of a clonally expanding tumor cell population. If no more than 10% of the excised tissue analyzed was non-tumor tissue, then for the ACB-PCR measured mutant fractions to be attributable to this cell fraction, the mutant fraction of each sample of non-tumor tissue would be ten times larger. This means that the mutant fractions in the non-tumor tissue would have to range from 0.5 to 2.3 x 10-4. Even in a chronically treated animal, it is very unlikely that the mutant fraction at a single basepair would reach these levels 71% of the time (22). Thus, it can be concluded that the mutant fraction data obtained by ACB-PCR provides evidence that a prototypical early ras mutation also occurs as a secondary event during tumor progression.

The 21 different tumors analyzed in this study were induced in male and female mice of two different strains using a number of different treatments. Consequently, no relationships could be discerned between treatment and mutation detection. Furthermore, there were not sufficient data available to relate tumor size to H-ras mutant fraction. However, all of the tumors included in the study had been subject to a pathological examination. Therefore, it was possible to consider the detection of H-ras mutation with respect to tumor type (Table IIIGo). Twelve hepatocellular adenomas were analyzed by ACB-PCR and the H-ras mutation was detected in 11 of them. As a benign neoplasia, adenoma is considered a relatively early stage in tumor development. With respect to these adenomas, even though the H-ras mutation being measured is a secondary mutation, it is still being detected at a relatively early stage of tumor development. Two of the tumors characterized by ACB-PCR were hepatocellular carcinomas, both of which had measurable H-ras mutant fractions. Carcinoma is a later, malignant stage in tumor development. With regard to these two tumor samples, therefore, the H-ras mutation was detected as a secondary event at a relatively late stage in tumor development. While H-ras mutation was detected in almost all the hepatocellular adenomas and carcinomas analyzed, it was detected in only one of five histiocytic sarcomas that were analyzed (20%, Table IIIGo). The same pattern of H-ras mutation detection with respect to tumor type was observed in the mutations initially detected by DGGE in Study #2 (previously unpublished observations from Culp et al. (17), Table IIIGo). The most likely explanation for this finding is that ras activation in different cell lineages has different phenotypic consequences. Depending upon the cell-type specific background, ras activation can lead to cell proliferation, cell arrest, or even apoptosis (23).

Several issues relevant to the measurement of oncomutations as small subpopulations are highlighted by this study. The characterization of tumors in Studies #1 and #2 identified tumors that did or did not have detectable H-ras mutation; the specific H-ras mutant fractions of the tumors were not determined. If mutations M5 and M6 (from Study #1) had been analyzed by DGGE in Study #2, most likely they would have been identified and reported as positive for H-ras mutation. It would be incorrect to assume that they were activating mutations. Clearly, the detection of tumors `positive' for a particular mutation should always be interpreted with regard to the sensitivity of the mutation detection method used. Activating oncogene mutations detected in tumors are not necessarily the tumor initiating mutations. The use of a quantitative assay, like ACB-PCR, makes this apparent. What then is the significance of mutations present at such low mutant fractions in terms of tumor development, progression, and metastasis? The literature shows that ras mutations can be detected at virtually every stage of carcinogenesis. The progression of carcinogenesis is thought to proceed through the accumulation of a number of different genetic changes, each of which confers a new tumorigenic phenotype on the cells acquiring them. Many different oncogenic lesions have been identified, each of which will have a certain finite probability or frequency of occurring. These are all relatively rare events and they can accumulate in different orders. This is why the accumulation of oncogenic lesions is a stochastic process. In the present study, an adenoma and carcinoma were each found to have H-ras mutant fractions of 3 x 10-5. One interpretation of this is that the carcinoma had acquired a greater number of genetic changes before the ras mutation occurred than the adenoma, but that they had approximately equal amounts of clonal amplification after acquiring their respective ras mutations (a mutant fraction of 3 x 10-5 in this experiment corresponds to the detection of approximately ten mutant cells).

This study suggests that the ACB-PCR could be used to characterize the accumulation of H-ras point mutations during tumor progression. However, the applicability of ACB-PCR to the study of tumor progression using other mutational targets will depend in large part upon the mutational sensitivity of each particular target. It should be noted that with the possible exception of K-ras codon 12 GGT to GAT mutation in pancreatic cancer, no single tumor-associated mutation is detected with such predominance in human or rodent tumors as the H-ras codon 61 CAA to AAA mutation is detected in mouse liver tumors (22). More work is needed to determine how many other specific oncogene or tumor suppressor gene mutations will occur with such a predominance during tumor progression that they could be analyzed using this approach.


    Notes
 
3 To whom correspondence should be addressed Email: bparsons{at}nctr.fda.gov Back


    Acknowledgments
 
The authors wish to acknowledge Edward Lew, Eddie E.Li, Peter P.Fu, Beverly A.Smith, and Frederick A.Beland for their contributions to this study.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

  1. Kinzler,K.W. and Vogelstein,B. (1996) Lessons from hereditary colorectal cancer. Cell, 87, 159–170.[ISI][Medline]
  2. D'Errico,M., Calcagnile,A. and Dogliotti,E. (1996) Genetic alterations in skin cancer. Ann. Istit. Sup. Sanit., 32, 53–63.
  3. Sekido,Y., Fong,K.M. and Minna,J.D. (1998) Progress in understanding the molecular pathogenesis of human lung cancer. Biochim. Biophys. Acta, 1378, F21–59.[ISI][Medline]
  4. Yokota,J. (2000) Tumor progression and metastasis. Carcinogenesis, 21, 497–503.[Abstract/Free Full Text]
  5. Hanahan,D. and Weinberg,R.A. (2000) The hallmarks of cancer. Cell, 100, 57–70.[ISI][Medline]
  6. Herrero-Jimenez,P., Thilly,G., Southam,P.J., Tomita-Mitchell,A., Morgenthaler,S., Furth,E.E. and Thilly,W.G. (1998) Mutation, cell kinetics and subpopulations at risk for colon cancer in the United States. Mutat. Res., 400, 553–578.[ISI][Medline]
  7. Renan,M.J. (1993) How many mutations are required for tumorigenesis? Implications from human cancer data. Mol. Carcinogen., 7, 139–146.[ISI][Medline]
  8. Simons,J.W. (1999) Genetic, epigenetic, dysgenetic and non-genetic mechanisms in tumorigenesis. II. Further delineation of the rate limiting step. Anticancer Res., 19, 4781–4789.[ISI][Medline]
  9. Periano,C., Raichards,W.L. and Stevens,F.J. (1983) Multistage hepatocarcinogenesis. In Slaga,T.J. (ed.) Mechanisms of Tumor Promotion. CRC Press, Boca Raton, FL, vol. 1, pp. 1–53.
  10. Maronpot,R.R., Fox,T., Malarkey,D.E. and Goldsworthy,T.L. (1995) Mutations in the ras proto-oncogene: clues to etiology and molecular pathogenesis of mouse liver tumors. Toxicology, 101, 125–156.[ISI][Medline]
  11. Mills,N.E., Fishman,C.L., Rom,W.N., Dubin,N. and Jacobson,D.R. (1995) Increased prevalence of K-ras oncogene mutations in lung adenocarcinoma. Cancer Res., 55, 1444–1447.[Abstract]
  12. Parsons,B.L. and Heflich,R.H. (1997) Genotypic selection methods for the direct analysis of point mutations. Mutat. Res., 387, 97–121.[ISI][Medline]
  13. Orou,A., Fechner,B., Utermann,G. and Menzel,H.J. (1995) Allele-specific competitive blocker PCR: a one-step method with applicability to pool screening. Hum. Mutat., 6, 163–169.[ISI][Medline]
  14. Parsons,B.L. and Heflich,R.H. (1998) Detection of a mouse H-ras codon 61 mutation using a modified allele-specific competitive blocker PCR genotypic selection method. Mutagenesis, 13, 581–588.[Abstract]
  15. Parsons,B.L. and Heflich,R.H. (1997) Evaluation of MutS as a tool for direct measurement of point mutations in genomic DNA. Mutat. Res., 374, 277–285.[ISI][Medline]
  16. Manjanatha,M.G., Li,E.E., Fu,P.P. and Heflich,R.H. (1996) H- and K-ras mutational profiles in chemically induced liver tumors from B6C3F1 and CD-1 mice. J. Toxicol. Environ. Health, 47, 195–208.[ISI][Medline]
  17. Culp,S.J., Warbritton,A.R., Smith,B.A., Li,E.E. and Beland,F.A. (2000) DNA adduct measurements, cell proliferation and tumor mutation induction in relation to tumor formation in B6C3F1 mice fed coal tar or benzo[a]pyrene. Carcinogenesis, 21, 1433–1440.[Abstract/Free Full Text]
  18. Culp,S.J., Gaylor,D.W., Sheldon,W.G., Goldstein,L.S. and Beland,F.A. (1998) A comparison of the tumors induced by coal tar and benzo[a]pyrene in a 2-year bioassay. Carcinogenesis, 19, 117–124.[Abstract]
  19. Bos,J.L. (1988) The ras gene family and human carcinogenesis. Mutat. Res., 195, 255–271.[ISI][Medline]
  20. Bos,J.L. (1989) ras oncogenes in human cancer: a review. Cancer Res., 49, 4682–4689.[Abstract]
  21. Parsons,B.L. and Heflich,R.H. (1998) Detection of basepair substitution mutation at a frequency of 1 x 10–7 by combining two genotypic selection methods, MutEx enrichment and allele-specific competitive blocker PCR. Environ. Mol. Mutagen., 32, 200–211.[ISI][Medline]
  22. McKinzie,P.B., Delongchamp,R.R., Heflich,R.H. and Parsons,B.L. (2001) Prospects for applying genotypic selection of somatic oncomutation to chemical risk assessment. Mutat. Res., 489, 47–78.[ISI][Medline]
  23. Frame,S. and Balmain,A. (2000) Integration of positive and negative growth signals during ras pathway activation in vivo. Curr. Opin. Genet. Develop., 10, 106–113.[ISI][Medline]
Received September 28, 2001; revised February 13, 2002; accepted March 1, 2002.





This Article
Abstract
FREE Full Text (PDF)
Alert me when this article is cited
Alert me if a correction is posted
Services
Email this article to a friend
Similar articles in this journal
Similar articles in ISI Web of Science
Similar articles in PubMed
Alert me to new issues of the journal
Add to My Personal Archive
Download to citation manager
Search for citing articles in:
ISI Web of Science (2)
Request Permissions
Google Scholar
Articles by Parsons, B. L.
Articles by Heflich, R. H.
PubMed
PubMed Citation
Articles by Parsons, B. L.
Articles by Heflich, R. H.