ARTICLE

Frequency of UV-Inducible NRAS Mutations in Melanomas of Patients With Germline CDKN2A Mutations

Malihe Eskandarpour, Jamileh Hashemi, Lena Kanter, Ulrik Ringborg, Anton Platz, Johan Hansson

Affiliation of authors: Department of Oncology-Pathology, Cancer Center Karolinska, Karolinska Hospital and Karolinska Institute, Stockholm, Sweden.

Correspondence to: Johan Hansson, M.D., Ph.D., Department of Oncology-Pathology, Karolinska Hospital, S-171 76, Stockholm, Sweden (e-mail: Johan. Hansson{at}onkpat.ki.se).


    ABSTRACT
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
Background: Germline alterations in cyclin-dependent kinase inhibitor 2A (CDKN2A) are important genetic factors in familial predisposition to melanoma. Activating mutations of the NRAS proto-oncogene are among the most common somatic genetic alterations in cutaneous malignant melanomas. We investigated the occurrence of NRAS mutations in melanomas and dysplastic nevi in individuals with germline CDKN2A mutations. Methods: Genomic DNA was extracted from 39 biopsy samples (including primary melanomas, metastatic melanomas, and dysplastic nevi) from 25 patients in six Swedish families with a hereditary predisposition to melanoma who carried germline CDKN2A mutations. DNA was also extracted from 10 biopsy samples from patients with sporadic melanomas. NRAS was analyzed using polymerase chain reaction, single-strand conformation polymorphism analysis, and nucleotide sequence analysis. Differences in NRAS mutation frequency between hereditary and sporadic melanomas were analyzed by the chi-square test. All statistical tests were two-sided. Results: Activating mutations in NRAS codon 61, all of which were either CAA(Gln)-AAA(Lys) or CAA(Gln)-CGA(Arg) mutations, were found in 95% (20/21) of primary hereditary melanomas but in only 10% (1/10) of sporadic melanomas (P<.001). Multiple activating NRAS mutations were detected in tumor cells from different regions of individual primary melanomas in nine patients. Activating mutations that were detected in the primary melanomas of these patients were also retained in their metastases. NRAS mutations at sites other than codon 61 were also present in the primary melanomas, indicating genetic instability of this locus. NRAS codon 61 mutations were also detected in dysplastic nevi and in an in situ melanoma, suggesting a role for such mutations during early melanoma development. Conclusions: The high frequency of NRAS codon 61 mutations detected in these hereditary melanomas may be the result of a hypermutability phenotype associated with a hereditary predisposition for melanoma development in patients with germline CDKN2A mutations.



    INTRODUCTION
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
Cutaneous malignant melanoma is a malignancy of melanocytes, the incidence of which is increasing in white-skinned populations throughout the world (1). Although the most important known risk factors are sun exposure and genetic predisposition, the molecular genetic alterations during the development of this disease are not well characterized. Mutations in many different genes, such as NRAS (25), BRAF (68), and PTEN (9,10), and mutations and deletions of the cyclin-dependent kinase inhibitor 2A (CDKN2A) (1115) have all been implicated in the pathogenesis of malignant melanoma.

The ras family of small G-proteins with guanosine triphosphate (GTP) hydrolyzing activity, which are expressed ubiquitously in mammalian cells, includes three closely related proteins: H-ras, K-ras, and N-ras. The three proto-oncogenes that encode these proteins have similar genetic structures, with four exons and a 5' non-coding exon (i.e., exon Ø). Ras proteins are involved in the transduction of extracellular signals (elicited by activating surface receptors) and act as key components in the relaying of signals downstream through diverse pathways. They play this role by virtue of their GTPase activity, functioning as guanosine diphosphate (GDP)/GTP-regulated switches that cycle between an active GTP-bound state and an inactive GDP-bound state (16,17).

Tumor-associated mutant ras proteins have single amino acid substitutions primarily at residues 12, 13, and 61, and to some extent at residues 59 and 63, that can render ras insensitive to GTPase activating protein (GAP)-stimulated GTP hydrolysis (17,18). Hence, oncogenic mutants of ras are chronically activated proteins that stay in the GTP-bound state and therefore continue to signal, even in the absence of extracellular stimuli (17,19).

Activating RAS mutations are among the most commonly identified genetic alterations in human cancers (20). Activating RAS mutations have been shown to be oncogenic in a transgenic mouse model (21) that showed that melanoma tumors result from expression of oncogenically mutated RAS in melanocytes. Oncogenically mutated RAS genes have been detected in up to 35% of human tumors, although the frequency of mutations for different RAS oncogenes varies, depending on the tissue of origin (20). Several studies with large numbers of melanoma patients have confirmed that oncogenic activation of NRAS constitutes the predominant RAS alteration in cutaneous melanoma (2,4,22). NRAS mutations are most often found in melanomas in sun-exposed areas of the body, suggesting an association between UV irradiation and the induction of NRAS mutations (4,22). Irradiation experiments with cloned human NRAS sequences and in vivo experiments in mice have also confirmed a link between UV exposure and NRAS activation (23,24).

The pathogenic role of alterations in the CDKN2A tumor suppressor gene, which has a key role in the CDK4–cyclin D–retinoblastoma protein (Rb) pathway and in the regulation of the G1 checkpoint of the cell cycle, has also been addressed in experimental and clinical studies of melanoma (11,14,25,26). In human melanoma cell lines, the CDKN2A gene is frequently mutated or homozygously deleted (27,28). CDKN2A alterations have also been reported in studies of sporadic cutaneous melanomas (11,13,25).

Germline CDKN2A mutations occur in many patients with a hereditary predisposition to melanoma. Overall, in approximately half of all melanoma-prone families, the disease shows genetic linkage to 9p21, the chromosome arm where the CDKN2A gene is located, and approximately 40% of these families carry germline mutations in CDKN2A (15,29). However, CDKN2A germline mutations are rare in Swedish families with a melanoma predisposition; they have been found in less than 10% of analyzed Swedish families (30). Such Swedish melanoma-prone families are identified through a national program, and thus these families represent a population-based set of melanoma families. A similarly small proportion of families carrying a germline CDKN2A mutation were observed in a population-based study in Queensland, Australia (31). A CDKN2A mutation consisting of a 3-base-pair (bp) insertion leading to an extra arginine residue in codon 113 in exon 2 (113insArg) has been found in 17 Swedish families (32). Haplotype analysis showed that these families originated from a common ancestor, and the age of the founder mutation was estimated at approximately 2000 years (32). Thus, the large majority of Swedish melanoma families with germline CDKN2A mutations have the same founder mutation. Because both CDKN2A alterations and activating NRAS mutations are implicated as important and frequent alterations in cutaneous melanoma, it is of interest to study the role of somatic NRAS mutations in melanomas in individuals with germline CDKN2A mutations. Moreover, the association between UV exposure and codon 61 NRAS mutations further supports the importance of investigating these alterations in individuals with an inherited increased susceptibility to develop these sun-related tumors.

In the only report of somatic NRAS mutations in hereditary melanomas to date (33), NRAS codon 61 mutations in melanoma metastases from patients with hereditary melanoma and dysplastic nevus syndrome (DNS) were statistically significantly more frequent than NRAS codon 61 mutations from patients with sporadic melanomas. However, the patients in that study were not characterized with respect to germline CDKN2A mutation status. The focus of the present study, therefore, was on NRAS alterations in tumor samples obtained from members of Swedish families who had hereditary melanoma and who carried germline CDKN2A mutations. Our aim was to determine the frequency of NRAS alterations in these patients and to establish the time of their appearance during tumor progression to better define the role of NRAS alterations in melanoma development in individuals carrying germline CDKN2A mutations.


    PATIENTS AND METHODS
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
Patients and Biopsy Samples

Thirty-nine biopsy samples from 25 patients belonging to six families with hereditary malignant melanoma and germline CDKN2A mutations were analyzed. All patients carried germline CDKN2A mutations with functional importance, as demonstrated by nucleotide sequence analysis of exons 1 and 2 of the CDKN2A gene (30). Four families carried the 113insArg germline mutation in exon 2 of CDKN2A, and one family carried the Pro48Leu germline mutation in exon 1 of CDKN2A. Two members of the latter family also carried the common codon 148 alanine to threonine single-nucleotide polymorphism in CDKN2A (30), which is a variant that is considered to have no functional importance. In a sixth family, one member carried a germline 24-bp deletion of codons 62–69 of CDKN2A (34). This investigation was approved by the Ethics Committee of the Karolinska Institute, and all patients provided informed consent.

Of the total of 39 biopsy samples, 28 were malignant lesions (21 primary melanomas, five metastatic melanomas, one breast carcinoma, and one squamous cell skin cancer), seven were dysplastic nevi, two were common nevi, and two were normal skin samples. For comparative purposes, 10 sporadic primary melanomas diagnosed in patients without a known family history of melanomas were also studied for NRAS alterations. All biopsy samples were formalin-fixed, paraffin-embedded tissue blocks that we obtained from the archives of five different pathology departments at hospitals in the Stockholm and Gothenburg areas of Sweden. Biopsy samples underwent laser-capture microdissection (LCM) to obtain well-defined cell populations for DNA extraction.

DNA Extraction

Four serial sections (6 µm) were cut from each paraffin-embedded biopsy sample, and one of the inner sections was counterstained with hematoxylin and eosin as a guide for LCM. After deparaffinization in xylene, the sections were stained with hematoxylin to localize the tumor area and were dehydrated with increasing concentrations of ethanol (70%, 95%, 99%) followed by xylene. The sections were then microdissected with a PixCell laser-capture microscope (Arcturus Engineering, Mountain View, CA). The dissected sections (30–200 cells) were incubated in 10–50 µL (depending on the number of cells in the target area) of lysis buffer (proteinase K at 1 mg/mL [Sigma-Aldrich, Steinheim, Germany] and 1% Tween 20 in TE buffer [10 mM Tris–HCl, 1 mM EDTA; pH 8.0]) at 42 °C overnight. Proteinase K was inactivated by incubating the samples in lysis buffer at 95 °C for 10 minutes. DNA was then extracted and purified using the Wizard Genomic DNA Purification Kit (Promega, Madison, WI), according to the manufacturer’s instructions.

For use as positive controls, DNA was also extracted from the human HT1080 fibrosarcoma and 224 metastatic melanoma cell lines by freeze–thaw incubations and proteinase K treatment, followed by DNA purification as outlined above. The HT1080 cell line carries an AAA(Lys) substitution, and the 224 metastatic melanoma cell line carries a CGA(Arg) substitution at codon 61 of the NRAS gene.

Polymerase Chain Reaction Amplification

Polymerase chain reaction (PCR) amplification was carried out in a nested or hemi-nested fashion. In the first PCR of NRAS exons 1 and 2, DNA extracts from captured cells (4-µL aliquots) or from the positive control cell lines (5 ng) were amplified in 25 µL of standardized buffer containing 0.5 M KCl, 0.1 M Tris–HCl (pH 8.3), and 2.5 mM MgCl2, by using the same amplification protocol. DNA samples and controls were amplified for 45 cycles at denaturing temperatures of 96 °C for 45 seconds and 35 cycles at 93 °C for 45 seconds. For the second PCR, DNA samples and controls were amplified in 10 µL of standardized buffer for 20 cycles at denaturing temperatures of 93 °C for 45 seconds, 60 °C for 45 seconds, and 72 °C for 45 seconds, with inner primers and products (1 µL) from the first PCR being used as a template. Fig. 1Go shows the sequences of the inner primers used in the PCR reactions. For exon 1, inner primer pairs 13S/13A and 13S/13AN were used with annealing temperatures of 60 °C and 61 °C, respectively, in a hemi-nested PCR manner, and for exon 2, inner primer pairs 61A/61B and 61S/61AS were used with annealing temperatures of 58 °C and 60 °C, respectively, in a nested PCR manner (Fig. 1Go). The final PCR products were labeled by incorporation of 3 µCi of [{alpha}-32P]dCTP (deoxycytidine triphosphate) per reaction. For each DNA extract, two independent PCR reactions were performed to confirm reproducibility.



View larger version (31K):
[in this window]
[in a new window]
 
Fig. 1. Inner primers used for polymerase chain reaction amplification of exons 1 and 2 of the NRAS gene.

 
Single-Strand Conformation Polymorphism and Nucleotide Sequencing Analysis

Single-strand conformation polymorphism (SSCP) analysis was carried out as described by Mashiyama et al. (35). Briefly, PCR products underwent electrophoresis on non-denaturing 7.5% acrylamide gels with 10% glycerol at 18 °C and 40 W. Shifted bands were excised from the gels and re-amplified by inner primers, and the purified PCR products were eluted from the acrylamide bands using a QIAquick Gel Extraction Kit (Qiagen GmbH, Hilden, Germany), according to the manufacturer’s instructions. Nucleotide sequence analyses for both DNA strands were performed by using the BigDye Terminator Cycle Sequencing Kit (Applied Biosystems, Foster City, CA) according to the manufacturer’s instructions (using the inner PCR primers shown in Fig. 1Go), and by using an ABI PRISM 310 Genetic Analyzer (Applied Biosystems). One sequencing reaction was performed for each DNA strand.

Statistical Analysis

Statistical differences in NRAS mutation frequencies between familial and sporadic melanomas were determined with the chi-square test for two independent samples by using StatView software (SAS Institute, Cary, NC). All statistical tests were two-sided.


    RESULTS
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
We analyzed melanomas and dysplastic nevi from individuals with germline CDKN2A mutations and a hereditary predisposition for the development of melanoma for the presence of activating somatic NRAS mutations. NRAS mutations were detected in approximately 96% (25/26) of primary and metastatic melanoma tumor samples from patients with CDKN2A germline mutations (Table 1Go). All activating NRAS mutations were located exclusively at codon 61 of exon 2, and no mutations were detected in codons 12 or 13 of exon 1. The NRAS codon 61 mutations consisted of a substitution of AAA(Lys) or CGA(Arg) for CAA(Gln), and tumors frequently had both mutations.


View this table:
[in this window]
[in a new window]
 
Table 1. Summary of NRAS mutations in tumor samples from individuals with germline CDKN2A mutations*
 
Activating codon 61 NRAS mutations were detected in 95% (20/21) of the primary melanoma tumors (Table 1Go). Arg and Lys substitutions at codon 61 were observed with approximately the same frequency (Table 2Go). Of the 20 primary melanomas with NRAS mutations, 17 were located in intermittently sun-exposed areas (leg, trunk), whereas two (tumors 18 and 21) were in chronically exposed areas (neck, hand), and one (tumor 8) was in a rarely exposed area (sole). No NRAS mutations other than those at codon 61 were detected in the primary melanoma samples. However, in nine tumor samples, mutation analysis revealed more than one activating mutation. Moreover, the NRAS genotype sometimes differed between extracts of melanoma cells from separate regions of the tumor (Fig. 2Go and Table 3Go).


View this table:
[in this window]
[in a new window]
 
Table 2. Mutation analysis of NRAS in DNA extracts from different areas of primary malignant melanomas*
 


View larger version (99K):
[in this window]
[in a new window]
 
Fig. 2. Laser capture microdissection (LCM) and single-strand conformation polymorphism (SSCP) analysis of a primary superficial spreading melanoma (SSM) tumor. A) Microdissection of a deparaffinized hematoxylin-stained section of the tumor. Photographs 1–3 show three different regions of the vertical growth phase of the same tumor from which DNA was extracted (tumor 7, Table 2Go). The left panel shows the tumor before microdissection, the middle panel shows the tumor after microdissection, and the right panel shows the laser-captured cells from which DNA was extracted. B) SSCP analyses of polymerase chain reaction products derived from exon 2 of NRAS on a 7.5% non-denaturing acrylamide gel. Lanes 1–3, SSCP analyses of DNA extracts from the three corresponding regions of the tumor depicted in A. Mutation analyses revealed an NRAS codon 61 Lys mutation in extract 1, an NRAS codon 61 Arg mutation in extract 2, and both mutations in extract 3. All tumor DNA extracts had the wild-type (wt) NRAS allele.

 

View this table:
[in this window]
[in a new window]
 
Table 3. NRAS mutations in multiple DNA extracts from common and dysplastic nevi*
 
In contrast to the hereditary melanomas, mutation analysis of 10 sporadic primary melanomas from patients with no known history of melanoma showed that only 10% (1/10) of primary tumors had a detectable NRAS mutation (CGA[Arg] substitution) at codon 61. This difference in mutation frequency of NRAS mutations between hereditary melanomas in patients with germline CDKN2A mutations and sporadic melanomas in patients with no known history of melanoma was statistically significant (P<.001; {chi}2 = 22.60). This finding confirms other reports of relatively low frequencies of NRAS mutations in sporadic melanomas (24,33).

To determine the time of appearance of NRAS mutations during tumor development, we also performed mutational analysis on both dysplastic nevus samples (i.e., precursor lesions) and one in situ melanoma sample. NRAS mutations at codon 61 were detected in the in situ melanoma sample (Table 2Go) and in five of seven dysplastic nevi samples (Tables 1Go and 3Go). In the dysplastic nevi samples, we detected complex mutation patterns—that is, in each dysplastic nevus, at least two separate NRAS codon 61 activating mutations were detected, and those mutations (Lys and Arg substitutions) sometimes occurred in different nests of melanoma cells (i.e., different DNA extracts from different parts of the lesion). NRAS mutations outside codon 61 were also detected (Table 3Go).

We also analyzed two normal skin tissue biopsies and two common nevi from four different patients who carried a germline 113insArg alteration in CDKN2A. LCM was carried out on different areas of normal tissue (data not shown) and on cells from common nevus cell nests (Table 3Go). Interestingly, none of the nevus cell populations had any mutated NRAS alleles.

To evaluate whether clones of primary melanoma cells with NRAS mutations were capable of expanding during tumor progression, we also analyzed five metastases that originated from primary melanomas in four patients (Table 4Go). The same NRAS mutations that were present in the primary tumors were also detected in all metastases from these patients, indicating a clonal relationship between melanoma cells in primary and metastatic melanoma tumors. However, additional NRAS mutations at sites other than at codon 61 were also present in these metastases. For example, in one patient who had a primary melanoma with an NRAS codon 61 Arg substitution with loss of the NRAS wild-type allele, the Arg mutation was retained in a lymph node metastasis derived from the primary tumor (Table 4Go). Four years later, the same patient had a new lymph node metastasis at the same site as the previous metastasis; however, this time, the tumor cells exhibited both the initial NRAS codon 61 Arg mutation and an Lys mutation, thus indicating that a melanoma cell clone carrying both mutations had expanded during tumor progression.


View this table:
[in this window]
[in a new window]
 
Table 4. NRAS mutations in primary malignant melanomas and their corresponding metastases
 
Twenty-two melanoma samples were heterozygous for the mutant NRAS allele, as indicated by the relative intensities of the SSCP bands representing the PCR products (Fig. 3Go, lane B1). However, in four melanoma tumor samples (tumors 9, 11, and 13 in Table 2Go and one metastatic tumor in Table 4Go), in which analyses were performed on homogenous tumor cell populations, the NRAS wild-type bands were completely lost, indicating loss of the NRAS wild-type allele (Tables 2Go and 4Go and Fig. 3Go). Interestingly, analysis of DNA extracts from different parts of the tumors showed different intensities of the shifted SSCP bands (Fig. 3Go), consistent with different proportions of tumor cells with mutant NRAS alleles in separate parts of the tumors.



View larger version (27K):
[in this window]
[in a new window]
 
Fig. 3. Single-strand conformation polymorphism (SSCP) analyses of polymerase chain reaction products derived from exon 2 of NRAS. A) NRAS mutation patterns in dysplastic nevi. Lanes 1 and 2, positive controls for NRAS codon 61 mutations. The HT1080 fibrosarcoma cell line, which is heterozygous for an AAA(Lys) substitution, is shown in lane 1, and the 224 melanoma cell line, which is hemizygous for a CGA(Arg) substitution, is shown in lane 2. A wild-type (wt) NRAS allele was also detected in lane 1. Lanes 3 and 4, DNA extracts from different regions of the same dysplastic nevus showing both activating NRAS codon 61 Lys and Arg mutations. Wild-type NRAS alleles were detected in both lanes. Note that the DNA mutation bands have different intensities, indicating an imbalance between the relative amounts of mutant alleles. Lane 5, loss of the wild-type NRAS allele in a dysplastic nevus that has an NRAS codon 61 Lys mutation, a codon 61 Arg mutation, and a codon 61 Arg mutation combined with an Arg + GAA62GAG (Glu-Glu) silent mutation. Lane 6, a dysplastic nevus showing a GAG63GGG (Glu-Gly) and an NRAS codon 61 Lys mutation. A wild-type NRAS allele was also detected. B) NRAS mutations from a primary melanoma and a corresponding metastasis from the same patient. Lanes 1 and 2, analysis of two DNA extracts from different regions of the primary melanoma, one of which has an NRAS codon 61 Lys mutation (lane 1) and a wild-type NRAS allele; the other has only a wild-type NRAS allele (lane 2). Lane 3, DNA extract from the metastasis shows four separate bands corresponding to an NRAS codon 61 Lys mutation, an Arg + GAA62GAG (Glu-Glu) mutation (indicated by *), a GAA63GAG (Glu-Glu) mutation (indicated by {dagger}), and a wild-type NRAS allele.

 
As expected, the NRAS codon 61 mutations were detected only in primary skin tumors, supporting the role of UV irradiation in the induction of such mutations. One of the patients with both a nodular and a superficial spreading melanoma also developed breast cancer (Tables 1Go and 2Go). The superficial spreading melanoma tumor from that patient had an NRAS codon 61 Lys mutation (AAA) (Table 2Go, tumor 15), and the nodular melanoma tumor had both Lys and Arg NRAS codon 61 mutations (AAA and CGA) (Table 2Go, tumor 14), whereas the breast tumor did not have any detectable NRAS mutations (Table 1Go). The two NRAS codon 61 mutations in the nodular melanoma tumor of that patient were detected in separate tumor cell extracts (Fig. 4Go). In contrast, a squamous cell carcinoma of the skin that occurred in a patient who had a germline CDKN2A mutation but who did not have a melanoma had both Lys and Arg NRAS codon 61 mutations (data not shown), which is consistent with the probable UV-associated etiology of this type of skin cancer.



View larger version (82K):
[in this window]
[in a new window]
 
Fig. 4. Laser capture microdissection (LCM) and single-strand conformation polymorphism (SSCP) analyses of a primary nodular melanoma tumor. A) Photographs 1 and 2 show two different regions of the tumor from which DNA was extracted (tumor 14, Table 2Go). The left panel shows the tumor before microdissection, the middle panel shows the tumor after microdissection, and the right panel shows the laser-captured cells from which DNA was extracted. B) SSCP analyses of polymerase chain reaction products derived from exon 2 of NRAS on a 7.5% non-denaturing acrylamide gel. Lanes 1 and 2 illustrate SSCP analyses of DNA extracts from the two corresponding regions of the tumor depicted in A. Mutation analyses revealed an NRAS codon 61 Lys mutation in extract 1 (lane 1) and an NRAS codon 61 Arg mutation in extract 2 (lane 2). Both DNA extracts also had the wild-type (wt) NRAS allele.

 

    DISCUSSION
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
This is the first report, to our knowledge, that documents a high frequency of NRAS mutations in primary melanomas in members of melanoma-prone families who also have known germline CDKN2A mutations. Our results support the concept of cooperation between CDKN2A inactivation and ras activation in the pathogenesis of human cutaneous melanoma.

We found that all biopsy samples with NRAS mutations exhibited activating codon 61 mutations, which are compatible with known mechanisms of UV mutagenesis. Van der Lubbe et al. (24) have shown that UV irradiation of cloned NRAS sequences followed by transfection into rat fibroblasts resulted in the appearance of transformed cells carrying vectors with the same NRAS codon 61 mutations that we found in our mutation analyses. The induction of CAA(Gln) to CGA(Arg) mutations by UV is not unexpected because the non-coding strand of codon 61 in NRAS contains a TT sequence, which is a classic site for UV-induced photo-products (24). Moreover, the first nucleotide of the non-coding strand in codon 61 of NRAS is a guanine base residue, which can be transformed to 8-oxo-guanine by UV irradiation (36). This modified base residue can then form a pair with an adenine residue during DNA replication, resulting in a CAA(Gln) to AAA(Lys) mutation.

By combining the techniques of LCM, PCR, SSCP, and DNA sequencing, we were able to detect mutations in very small amounts of template DNA. A possible source of artifacts in this study is the induction of DNA replication errors during PCR amplification. Because such replication errors do occur, we took precautions to prevent such artifacts by performing two independent PCR reactions on each DNA extract and using the results only when both SSCP analyses were identical. In addition, activating NRAS codon 61 mutations, which are our predominant findings in this study, have been previously reported in melanoma tumors by several investigators (24,22,33) and are therefore extremely unlikely to be the result of PCR errors. Moreover, we always used parallel PCR reactions without DNA template as negative controls, and no amplification products were obtained in those control PCR reactions, indicating that contamination of DNA from other sources did not occur.

The LCM technique enabled us to investigate the issue of clonal heterogeneity among melanoma cell clones in individual tumors. Thus, in several cases, we were able to analyze DNA extracts from multiple regions of the same primary melanoma. Different mutation patterns were seen in separate regions of individual tumors, which may reflect the expansion of several different tumor cell clones during melanoma development (Figs. 2Go and 4Go). In some tumors, SSCP analyses detected a loss of the wild-type NRAS allele in the tumor cells. Recent observations in animal models (37) suggest that the loss of the wild-type RAS allele may facilitate tumorigenesis induced by the remaining oncogenically mutated RAS allele. In addition to the loss of the wild-type NRAS allele, we also detected mutational activation of both NRAS alleles in some melanoma tumors, indicating genetic instability that may enhance the development of a malignant tumor cell phenotype.

NRAS codon 61 mutations have been reported to be statistically significantly more common in sun-exposed skin areas than in non-exposed mucosal membranes (4,22,38). Consistent with this notion, all but one of the primary melanomas in our study were located in chronically or intermittently sun-exposed areas of the body. In addition, an activating NRAS codon 61 mutation was also detected in a squamous cell carcinoma of the skin in one of the patients, whereas no mutation was detected in a breast carcinoma in another patient, further supporting the role of UV irradiation in the induction of NRAS codon 61 mutations. Therefore, our data provide molecular evidence for the importance of UV mutagenesis in the development of melanomas in melanoma-prone individuals with CDKN2A germline mutations.

NRAS mutations were also frequently present in dysplastic nevi and in the one in situ melanoma tumor that we analyzed, suggesting that NRAS mutations may be early events in the development of melanoma in individuals with germline CDKN2A mutations (Fig. 2Go). In contrast, no NRAS mutations were detected in common nevi and normal skin tissue biopsies. In a recent study of sporadic melanomas (5), we also found that NRAS codon 61 mutations in the radial growth phase of melanomas and nevi were associated with primary melanoma tumors, which supports the theory that NRAS codon 61 mutations are early events in the development of melanomas. In this respect, our finding that NRAS codon 61 mutations are early events in the development of both sporadic and hereditary melanomas may be somewhat different from the results of Demunter et al. (39,40), who reported that codon 61 NRAS mutations may occur both early and later during the progression of sporadic cutaneous melanomas. These differences in results may be affected by methodologic differences. The timing of NRAS mutations during melanoma development may be further elucidated by careful mutation studies of melanomas and precursor lesions using microdissection.

At the molecular epidemiologic level, the role of a cooperative effect of germline CDKN2A mutations and UV irradiation in the development of melanoma has been further supported by a Melanoma Genetics Consortium study (41), which found that the penetrance of germline CDKN2A mutations with respect to the development of melanoma was dependent on the country of residence. Higher melanoma risk was found in individuals living in countries with high environmental UV exposure, such as Australia, than in individuals living in countries with low environmental UV exposure, such as those in Europe. Unfortunately, it was not possible to perform a meaningful penetrance analysis on Swedish melanoma-prone families with germline CDKN2A mutations because only six such families from the Stockholm area in Sweden were included in the Melanoma Genetics Consortium study (41). However, the available data on melanoma development in the Swedish families indicate that the penetrance of CDKN2A mutations is higher among them than it is among families from other European countries and that it is similar to the penetrance estimates of CDKN2A mutations in families from the United States (Bishop T: personal communication).

Cooperation between inactivation of CDKN2A and activating mutations in RAS genes in melanoma development at the molecular level has been demonstrated in animal models. For example, Chin et al. (21) have provided in vivo experimental evidence for an association between CDKN2A deficiency and activating HRAS codon 12 (Gly-Val) mutations during the pathogenesis of melanoma by showing that melanocyte-specific expression of mutated HRAS in mice deficient for CDKN2A leads to the development of melanomas at a high frequency. This finding suggests a synergistic action of RAS activation and CDKN2A deficiency in melanocyte transformation. Thus, our finding of a high frequency of activating NRAS codon 61 mutations in melanomas in which the CDKN2A gene has been inactivated by a germline mutation supports the concept of CDKN2A inactivation and ras activation as key genetic alterations in human melanoma.

The high frequency of NRAS codon 61 mutations in this study is consistent with a UV hypermutability phenotype in patients with germline CDKN2A mutations and a hereditary predisposition to melanoma. Previous experimental studies (4245) have provided evidence that members of melanoma-prone families with dysplastic nevi may have a UV hypermutability phenotype. For example, Epstein–Barr virus-transformed lymphoblastoid cell lines from patients with hereditary dysplastic nevus syndrome were shown to be hypermutable to UV radiation (42). In other studies (4345), shuttle vector plasmid systems were used to demonstrate increased UV mutability in lymphoblastoid cell lines from members of melanoma-prone families with dysplastic nevus syndrome. The authors of those studies concluded that members of families with a hereditary predisposition for melanoma and dysplastic nevus syndrome have a defective mechanism for handling UV-induced DNA damage; however, the nature of such a defective mechanism has not been defined. It should be noted, however, that those studies were performed on cell lines from individuals who had not been characterized with respect to germline CDKN2A mutations. Our results, therefore, provide the first molecular genetic data on melanoma tumors, to our knowledge, that support the theory that members of melanoma-prone families with germline CDKN2A mutations exhibit a hypermutability phenotype and that this hypermutability is linked to the induction of melanomas in these families.

Although the underlying mechanism of UV hypermutability is unknown, it may be related to the germline CDKN2A mutations in high-risk melanoma-prone individuals and, therefore, to the role of the p16 protein in the G1 checkpoint control of the cell cycle (27,46). Because the majority of the patients investigated in our study carried the same Swedish CDKN2A 113insArg germline founder mutation (32), it would be of interest to extend NRAS mutation analyses to melanomas arising in members of melanoma-prone families that have a larger spectrum of germline CDKN2A mutations and to individuals belonging to families with hereditary melanoma and wild-type CDKN2A.

Our finding that NRAS mutations are maintained during tumor progression are consistent with results from animal models. Studies using a vector system with a doxycycline-inducible mutant codon 12 (Gly-Val) of HRAS in CDKN2A-deficient transgenic mice showed that expression of mutant ras is essential for tumor maintenance (21). The fact that activating NRAS mutations were not lost in any of the metastases that we studied supports the hypothesis that such mutations may be essential for tumor cell proliferation. The importance of activating NRAS codon 61 mutations in tumor cell proliferation was further supported in a recent study of sporadic melanomas (5), in which NRAS codon 61 mutations identical to those detected in the primary melanomas in this study were maintained in essentially all metastases investigated. Therefore, further studies that investigate the importance of mutationally activated NRAS in human melanoma, for example, of specifically targeting the mRNAs of the mutated NRAS gene with short interfering RNA (siRNA) (4749), are warranted. The finding that activating mutations in the BRAF gene (one of the downstream targets of RAS genes) occur frequently in melanoma tumors (68) provides further evidence for the importance of the ras-raf-mitogen-activated protein kinase (MAPK) signal transduction pathway for melanoma development.

The accumulation of novel NRAS mutations in expanding tumor cell clones that also carried the original activating NRAS codon 61 mutations (Fig. 3Go) provides support for the notion of genetic instability in high-risk individuals with germline CDKN2A mutations. Interestingly, although absent in primary melanomas, additional NRAS mutations other than at codon 61 were also detected in some dysplastic nevi. The reason for this difference in mutation patterns between dysplastic nevi and primary melanomas remains unclear, but one possibility is that additional NRAS mutations can accumulate over time in the quiescent nevus cell populations of dysplastic nevi but may be lost from actively proliferating tumor cell clones in primary melanomas. Although the normally observed activating NRAS codon 61 mutations are UV inducible, several of these additional NRAS mutations are not compatible with known mechanisms of UV mutagenesis. In addition, although certain NRAS mutations, such as the codon 63 Glu to Gly substitution observed in dysplastic nevi, activate NRAS (17,18), these mutations are frequently located at sites where mutations have no known functional effects and where some of them are silent. Therefore, these additional NRAS mutations may not have a functional role in tumor progression but may reflect a general hypermutability phenotype in the dysplastic nevi of individuals with germline CDKN2A mutations.

In summary, our analysis of NRAS mutations in melanomas and dysplastic nevi in individuals with germline CDKN2A mutations provides new information in regard to key events in the development of melanoma in high-risk individuals and highlights the cooperative relationship between environmental UV exposure and inherited germline mutations in carcinogenesis. Our data indicate that members of melanoma-prone families with germline CDKN2A mutations exhibit a UV hypermutability phenotype that may lead to the frequent occurrence of UV-induced mutations in melanoma precursor lesions, such as dysplastic nevi. The combination of germline inactivation of the tumor suppressor gene CDKN2A and somatic activating mutations in the NRAS proto-oncogene are likely to be key genetic alterations contributing to the increased risk of developing melanoma from precursor lesions in these high-risk individuals. The importance of this combination of genetic alterations in melanoma tumor progression is supported by the finding that the original activating NRAS mutations are also present in melanoma metastases from the same patients. In addition, the accumulation of additional NRAS mutations other than at codon 61 in dysplastic nevi and melanoma metastases, which are not normally seen in sporadic melanoma tumors, may reflect a general genetic instability in these high-risk melanoma-prone individuals.

The results also suggest that activating NRAS may be an important target for specific anticancer therapy in melanoma patients who carry codon 61 mutations. Patients with hereditary melanoma and germline CDKN2A mutations, whose tumors generally have activating NRAS mutations, represent an appropriate candidate group for the development of such therapies. Moreover, because these mutations appear to occur early in tumor development, the concept of using NRAS codon 61 mutations as a target for the development of specific interventions might also be extended to include future trials of specific methods of prevention. At present, oncogenically activated NRAS genes may be targeted by farnesyl transferase inhibitors and by immunotherapy using specific peptide vaccines directed against NRAS codon 61 mutants (19,5053). Further studies of the gene expression profile in melanoma tumors with activating NRAS mutations may lead to the identification of novel targets for preventive and therapeutic strategies in melanoma-prone individuals.


    NOTES
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 
Supported by the Cancer Society of Stockholm, the King Gustav V Jubilee Fund, the Karolinska Institute Research Funds, and the Swedish Radiation Protection Institute.

We thank Milan Ridderikhof for technical assistance and Margareta Rodensjö for assistance with preparation of microscopy slides.


    REFERENCES
 Top
 Notes
 Abstract
 Introduction
 Patients and Methods
 Results
 Discussion
 References
 

1 Berwick M. Epidemiology. Current trends, risk factors, and environmental concerns. In: Balch CM, Houghton AN, Sober AJ, Soong S-j, editors. Cutaneous melanoma. 3rd ed. St. Louis (MO): Quality Medical Publishing; 1998. p. 551–71.

2 Ball NJ, Yohn JJ, Morelli JG, Norris DA, Golitz LE, Hoeffler JP. Ras mutations in human melanoma: a marker of malignant progression. J Invest Dermatol 1994;102:285–90.[Abstract]

3 Jafari M, Papp T, Kirchner S, Diener U, Henschler D, Burg G, et al. Analysis of ras mutations in human melanocytic lesions: activation of the ras gene seems to be associated with the nodular type of human malignant melanoma. J Cancer Res Clin Oncol 1995;121:23–30.[ISI][Medline]

4 Jiveskog S, Ragnarsson-Olding B, Platz A, Ringborg U. N-ras mutations are common in melanomas from sun-exposed skin of humans but rare in mucosal membranes or unexposed skin. J Invest Dermatol 1998;111:757–61.[Abstract]

5 Omholt K, Karsberg S, Platz A, Kanter L, Ringborg U, Hansson J. Screening of N-ras codon 61 mutations in paired primary and metastatic cutaneous melanomas: mutations occur early and persist throughout tumor progression. Clin Cancer Res 2002;8:3468–74.[Abstract/Free Full Text]

6 Davies H, Bignell GR, Cox C, Stephens P, Edkins S, Clegg S, et al. Mutations of the BRAF gene in human cancer. Nature 2002;417: 949–54.[CrossRef][ISI][Medline]

7 Brose MS, Volpe P, Feldman M, Kumar M, Rishi I, Gerrero R, et al. BRAF and RAS mutations in human lung cancer and melanoma. Cancer Res 2002;62:6997–7000.[Abstract/Free Full Text]

8 Pollock PM, Harper UL, Hansen KS, Yudt LM, Stark M, Robbins CM, et al. High frequency of BRAF mutations in nevi. Nat Genet 2003;33:19–20.[CrossRef][ISI][Medline]

9 Birck A, Ahrenkiel V, Zeuthen J, Hou-Jensen K, Guldberg P. Mutation and allelic loss of the PTEN/MMAC1 gene in primary and metastatic melanoma biopsies. J Invest Dermatol 2000;114:277–80.[Abstract/Free Full Text]

10 Guldberg P, thor Straten P, Birck A, Ahrenkiel V, Kirkin AF, Zeuthen J. Disruption of the MMAC1/PTEN gene by deletion or mutation is a frequent event in malignant melanoma. Cancer Res 1997;57:3660–3.[Abstract]

11 Kumar R, Lundh Rozell B, Louhelainen J, Hemminki K. Mutations in the CDKN2A (p16INK4a) gene in microdissected sporadic primary melanomas. Int J Cancer 1998;75:193–8.[CrossRef][ISI][Medline]

12 Kumar R, Smeds J, Lundh Rozell B, Hemminki K. Loss of heterozygosity at chromosome 9p21 (INK4-p14ARF locus): homozygous deletions and mutations in the p16 and p14ARF genes in sporadic primary melanomas. Melanoma Res 1999;9:138–47.[ISI][Medline]

13 Piccinin S, Doglioni C, Maestro R, Vukosavljevic T, Gasparotto D, D’Orazi C, et al. p16/CDKN2 and CDK4 gene mutations in sporadic melanoma development and progression. Int J Cancer 1997;74:26–30.[CrossRef][ISI][Medline]

14 Hussussian CJ, Struewing JP, Goldstein AM, Higgins PA, Ally DS, Sheahan MD, et al. Germline p16 mutations in familial melanoma. Nat Genet 1994;8:15–21.[ISI][Medline]

15 Platz A, Ringborg U, Hansson J. Hereditary cutaneous melanoma. Semin Cancer Biol 2000;10:319–26.[CrossRef][ISI][Medline]

16 Pruitt K, Der CJ. Ras and Rho regulation of the cell cycle and oncogenesis. Cancer Lett 2001;171:1–10.[CrossRef][ISI][Medline]

17 Crespo P, Leon J. Ras proteins in the control of the cell cycle and cell differentiation. Cell Mol Life Sci 2000;57:1613–36.[ISI][Medline]

18 Lowy DR, Willumsen BM. Function and regulation of ras. Annu Rev Biochem 1993;62:851–91.[CrossRef][ISI][Medline]

19 Adjei AA. Blocking oncogenic Ras signaling for cancer therapy. J Natl Cancer Inst 2001;93:1062–74.[Abstract/Free Full Text]

20 Macaluso M, Russo G, Cinti C, Bazan V, Gebbia N, Russo A. Ras family genes: an interesting link between cell cycle and cancer. J Cell Physiol 2002;192:125–30.[CrossRef][ISI][Medline]

21 Chin L, Pomerantz J, Polsky D, Jacobson M, Cohen C, Cordon-Cardo C, et al. Cooperative effects of INK4a and ras in melanoma susceptibility in vivo. Genes Dev 1997;11:2822–34.[Abstract/Free Full Text]

22 van Elsas A, Zerp SF, van der Flier S, Kruse KM, Aarnoudse C, Hayward NK, et al. Relevance of ultraviolet-induced N-ras oncogene point mutations in development of primary human cutaneous melanoma. Am J Pathol 1996;149:883–93.[Abstract]

23 Pierceall WE, Kripke ML, Ananthaswamy HN. N-ras mutation in ultraviolet radiation-induced murine skin cancers. Cancer Res 1992;52:3946–51.[Abstract]

24 Van der Lubbe JL, Rosdorff HJ, Bos JL, Van der Eb AJ. Activation of N-ras induced by ultraviolet irradiation in vitro. Oncogene Res 1988;3:9–20.[ISI][Medline]

25 Platz A, Ringborg U, Lagerlof B, Lundqvist E, Sevigny P, Inganas M. Mutational analysis of the CDKN2 gene in metastases from patients with cutaneous malignant melanoma. Br J Cancer 1996;73:344–8.[ISI][Medline]

26 Kamb A, Shattuck-Eidens D, Eeles R, Liu Q, Gruis NA, Ding W, et al. Analysis of the p16 gene (CDKN2) as a candidate for the chromosome 9p melanoma susceptibility locus. Nat Genet 1994;8:23–6.[Medline]

27 Sherr CJ. Cancer cell cycles. Science 1996;274:1672–7.[Abstract/Free Full Text]

28 Castellano M, Pollock PM, Walters MK, Sparrow LE, Down LM, Gabrielli BG, et al. CDKN2A/p16 is inactivated in most melanoma cell lines. Cancer Res 1997;57:4868–75.[Abstract]

29 Haluska FG, Hodi FS. Molecular genetics of familial cutaneous melanoma. J Clin Oncol 1998;16:670–82.[Abstract]

30 Platz A, Hansson J, Mansson-Brahme E, Lagerlof B, Linder S, Lundqvist E, et al. Screening of germline mutations in the CDKN2A and CDKN2B genes in Swedish families with hereditary cutaneous melanoma. J Natl Cancer Inst 1997;89:697–702.[Abstract/Free Full Text]

31 Aitken J, Welch J, Duffy D, Milligan A, Green A, Martin N, et al. CDKN2A variants in a population-based sample of Queensland families with melanoma. J Natl Cancer Inst 1999;91:446–52.[Abstract/Free Full Text]

32 Hashemi J, Bendahl PO, Sandberg T, Platz A, Linder S, Stierner U, et al. Haplotype analysis and age estimation of the 113insR CDKN2A founder mutation in Swedish melanoma families. Genes Chromosomes Cancer 2001;31:107–16.[CrossRef][ISI][Medline]

33 Platz A, Ringborg U, Brahme EM, Lagerlof B. Melanoma metastases from patients with hereditary cutaneous malignant melanoma contain a high frequency of N-ras activating mutations. Melanoma Res 1994;4:169–77.[ISI][Medline]

34 Hashemi J, Platz A, Ueno T, Stierner U, Ringborg U, Hansson J. CDKN2A germ-line mutations in individuals with multiple cutaneous melanomas. Cancer Res 2000;60:6864–7.[Abstract/Free Full Text]

35 Mashiyama S, Murakami Y, Yoshimoto T, Sekiya T, Hayashi K. Detection of p53 gene mutations in human brain tumors by single-strand conformation polymorphism analysis of polymerase chain reaction products. Oncogene 1991;6:1313–8.[ISI][Medline]

36 Epe B. DNA damage profiles induced by oxidizing agents. Rev Physiol Biochem Pharmacol 1996;127:223–49.[ISI][Medline]

37 Zhang Z, Wang Y, Vikis HG, Johnson L, Liu G, Li J, et al. Wildtype Kras2 can inhibit lung carcinogenesis in mice. Nat Genet 2001;29:25–33.[CrossRef][ISI][Medline]

38 van Elsas A, Scheibenbogen C, van der Minne C, Zerp SF, Keilholz U, Schrier PI. UV-induced N-ras mutations are T-cell targets in human melanoma. Melanoma Res 1997;7 Suppl 2:S107–13.[ISI][Medline]

39 Demunter A, Ahmadian MR, Libbrecht L, Stas M, Baens M, Scheffzek K, et al. A novel N-ras mutation in malignant melanoma is associated with excellent prognosis. Cancer Res 2001;61:4916–22.[Abstract/Free Full Text]

40 Demunter A, Stas M, Degreef H, De Wolf-Peeters C, van den Oord JJ. Analysis of N- and K-ras mutations in the distinctive tumor progression phases of melanoma. J Invest Dermatol 2001;117:1483–9.[Abstract/Free Full Text]

41 Bishop DT, Demenais F, Goldstein AM, Bergman W, Bishop JN, Paillerets BB, et al. Geographical variation in the penetrance of CDKN2A mutations for melanoma. J Natl Cancer Inst 2002;94:894–903.[Abstract/Free Full Text]

42 Perera MI, Um KI, Greene MH, Waters HL, Bredberg A, Kraemer KH. Hereditary dysplastic nevus syndrome: lymphoid cell ultraviolet hypermutability in association with increased melanoma susceptibility. Cancer Res 1986;46:1005–9.[Abstract]

43 Seetharam S, Waters HL, Seidman MM, Kraemer KH. Ultraviolet mutagenesis in a plasmid vector replicated in lymphoid cells from patient with the melanoma-prone disorder dysplastic nevus syndrome. Cancer Res 1989;49:5918–21.[Abstract]

44 Moriwaki S, Tarone RE, Kraemer KH. A potential laboratory test for dysplastic nevus syndrome: ultraviolet hypermutability of a shuttle vector plasmid. J Invest Dermatol 1994;103:7–12.[Abstract]

45 Moriwaki SI, Tarone RE, Tucker MA, Goldstein AM, Kraemer KH. Hypermutability of UV-treated plasmids in dysplastic nevus/familial melanoma cell lines. Cancer Res 1997;57:4637–41.[Abstract]

46 Shapiro GI, Edwards CD, Rollins BJ. The physiology of p16(INK4A)-mediated G1 proliferative arrest. Cell Biochem Biophys 2000;33:189–97.[CrossRef][ISI][Medline]

47 Bernstein E, Denli AM, Hannon GJ. The rest is silence. RNA 2001;7:1509–21.[Abstract/Free Full Text]

48 Paddison PJ, Caudy AA, Bernstein E, Hannon GJ, Conklin DS. Short hairpin RNAs (shRNAs) induce sequence-specific silencing in mammalian cells. Genes Dev 2002;16:948–58.[Abstract/Free Full Text]

49 Paddison PJ, Caudy AA, Hannon GJ. Stable suppression of gene expression by RNAi in mammalian cells. Proc Natl Acad Sci U S A 2002;99:1443–8.[Abstract/Free Full Text]

50 Abrams SI, Hand PH, Tsang KY, Schlom J. Mutant ras epitopes as targets for cancer vaccines. Semin Oncol 1996;23:118–34.[ISI][Medline]

51 Hunger RE, Brand CU, Streit M, Eriksen JA, Gjertsen MK, Saeterdal I, et al. Successful induction of immune responses against mutant ras in melanoma patients using intradermal injection of peptides and GM-CSF as adjuvant. Exp Dermatol 2001;10:161–7.[CrossRef][ISI][Medline]

52 Sebti SM, Hamilton AD. Farnesyltransferase and geranylgeranyltransferase I inhibitors in cancer therapy: important mechanistic and bench to bedside issues. Expert Opin Investig Drugs 2000;9:2767–82.[ISI][Medline]

53 Haluska P, Dy GK, Adjei AA. Farnesyl transferase inhibitors as anticancer agents. Eur J Cancer 2002;38:1685–700.[CrossRef][ISI][Medline]

54 Clark WH Jr, From L, Bernardino EA, Mihm MC. The histogenesis and biologic behavior of primary human malignant melanomas of the skin. Cancer Res 1969;29:705–27.[ISI][Medline]

Manuscript received July 21, 2002; revised March 27, 2003; accepted April 8, 2003.


This article has been cited by other articles in HighWire Press-hosted journals:


             
Copyright © 2003 Oxford University Press (unless otherwise stated)
Oxford University Press Privacy Policy and Legal Statement