Affiliations of authors: Q. Wei, S. S. Strom, L. Wang, Z. Guo, Y. Qiao, C. I. Amos, M. R. Spitz (Department of Epidemiology), J. E. Lee, J. E. Gershenwald, M. I. Ross, P. F. Mansfield (Department of Surgical Oncology), M. Duvic (Department of Dermatology), The University of Texas M. D. Anderson Cancer Center, Houston.
Correspondence to: Qingyi Wei, M.D., Ph.D., Department of Epidemiology, Unit 189, The University of Texas M. D. Anderson Cancer Center, 1515 Holcombe Blvd., Houston, TX 77030 (e-mail: qwei{at}mdanderson.org).
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
It is well documented that sunlight exposure is directly associated with CMM in humans (7). Intermittent sun exposure early in life, rather than cumulative sun exposure, appears to be the major risk factor for CMM (8). Increased sensitivity to the acute effects of sunlight exposure (characterized by erythema and sunburns) is also associated with an increased risk of CMM (9). This association is further supported by the fact that childhood immigration to Australia, where the ambient UV exposure is the highest in the world, is associated with an increase in the lifetime risk of CMM (10). The risk of CMM increases with age and is also increased in individuals with light skin color, or those who freckle, sunburn, or do not tan (11), further suggesting that sunlight exposure has a role in the etiology of CMM.
The fact that only a fraction of those exposed to sunlight develop CMM suggests that genetic susceptibility may have a role in the etiology of sunlight-induced CMM. For instance, those who had a strong family history of dysplastic nevus syndrome (12,13) were found to have an increased risk of CMM. However, common, benign, atypical, and dysplastic nevi associated with sunlight exposure are independent predictors of sporadic melanomas (14,15), suggesting that a low capacity for DNA repair may be the underlying molecular mechanism for sporadic melanomas. In familial atypical multiple-mole melanoma syndrome, inherited cutaneous moles or nevi appear to be the precursors of malignant lesions (16,17). Although some melanoma susceptibility genes, such as p16/MTS1/CDKN2A (18), are mutated in individuals with familial melanoma, hereditary melanoma accounts for only about 10% of all melanomas (19). Therefore, other susceptibility genes may contribute to the more than 90% of sporadic CMM that may be induced by sunlight exposure. There is a high frequency and early onset of melanomas in patients with xeroderma pigmentosum (XP), a rare (one in 250 000 individuals in the United States) autosomal recessive disease of mutated DNA repair genes resulting in marked UV hypersensitivity (20). This finding further suggests that genetically determined deficiencies in DNA repair capacity (DRC) may contribute to sunlight-induced CMM.
In a recent Italian casecontrol study of 132 CMM patients and 145 control subjects from a population with a high risk of familial dysplastic nevi, DRC was determined by a host-cell reactivation assay that measures nucleotide excision repair (21), but DRC was not found to be an independent risk factor for CMM (13). Because it is possible that these results would be different for a patient population with a high rate of sporadic disease, we used the same assay to retest the hypothesis that reduced DRC for UV light-induced DNA damage is associated with sporadic CMM.
![]() |
SUBJECTS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Case patients were recruited from The University of Texas M. D. Anderson Cancer Center (Houston, TX). All patients with either newly diagnosed or surgically treated, histopathologically confirmed CMM who were Texas residents and registered at the Cancer Center between April 1994 and July 2001 were eligible for our study. However, only untreated case patients (22% of the eligible cases) were recruited for this study. Cancer-free control subjects were recruited from among genetically unrelated clinic visitors (82.7%) and spouses (17.3%) and were individually matched to the case patients by age (±5 years), sex, and ethnicity. The exclusion criteria for case patients were prior chemotherapy or radiation therapy and metastasis; the exclusion criteria for all study subjects were prior cancer (except for nonmelanoma skin cancer for the case patients) and any blood transfusion in the 6 months prior to recruitment. The participation rate was greater than 90% among both case patients and control subjects. After we obtained written informed consent, each subject was given a structured self-administered questionnaire to collect demographic data and data on risk factors, such as natural hair color, eye color, skin color, presence of moles and dysplastic nevi, history of sunlight exposure (including freckling in the sun as a child, tanning ability, and number of sunburns), and of the number of first-degree relatives with any cancer. The study protocol was approved by the Institutional Review Board of The University of Texas M. D. Anderson Cancer Center.
Blood Processing and Lymphoblastoid Cell Lines
Each subject donated a 30-mL blood sample that was drawn into a heparinized Vacutainer (BD Biosciences, Franklin Lakes, NJ). The lymphocytes were isolated from each sample within 8 hours of blood collection by Ficoll-gradient centrifugation (21), resuspended, and frozen in freezing medium (50% fetal bovine serum, 40% RPMI-1640, 10% dimethyl sulfoxide), and stored at -80 °C. (No samples were stored for more than 30 months before being assayed). We used four Epstein-Barr virus-immortalized human lymphoblastoid cell lines from the Human Genetic Mutant Cell Repositories (Camden, NJ) as experimental controls. Two of the cell lines, GM00892B and GM00131A, have apparently normal DRCs of approximately 20% and 15%, respectively, as determined by the host-cell reactivation assay using plasmids damaged by 800 J/m2 UV. The other two cell lines were derived from XP patients and are deficient in nucleotide excision repair: GM02345B is an XP group A line with a DRC of less than 1%, and GM02246B is an XP group C line with a DRC of approximately 1%.
The Host-Cell Reactivation Assay
We used the host-cell reactivation assay to measure cellular DRC (21). This assay is based on the principle that if a reporter gene in a nonreplicating plasmid is damaged before the plasmid is transfected into cells, its expression in cells is dependent on the ability of the cells to repair the damage. We assumed that the DRC of the lymphocytes collected from the study subjects reflected the overall repair capacity of the donors, because in the DNA repair deficiency syndrome XP, deficient DRC is detected in all tissues, including lymphocytes (20). To measure the ability of cells to remove UV light-induced DNA damage (e.g., photoproducts) from a reporter gene encoding chloramphenicol acetyltransferase (CAT) in the plasmid pCMVcat (a gift from Lawrence Grossman of Johns Hopkins University, Baltimore, MD) (21), we transfected phytohemagglutinin-stimulated T lymphocytes and the four human lymphoblastoid cell lines with damaged and undamaged plasmids. The damaged CAT gene should not be expressed unless host-cell repair has occurred, because even one unrepaired photoproduct (such as a pyrimidine dimer) can effectively block expression of the reporter gene (22). Therefore, this assay provides a quantitative measurement of the nucleotide excision repair phenotype of the donor.
Lymphocyte samples were coded and run in batches of 1020 samples by personnel who were blinded to case or control status of the donors. Frozen cells were thawed and assayed for DRC as previously described (21). Briefly, the cells in each vial (1.5 mL) were quickly thawed and mixed before the last trace of ice disappeared with 7.0 mL of thawing medium (50% fetal bovine serum, 40% RPMI-1640, 10% dextrose), which ensured a cellular viability of more than 80%, as tested by exclusion of 0.4% trypan blue (Sigma Chemical Co., St. Louis, MO) (21). The cells were then washed with the thawing medium, resuspended at 106 cells per mL of RPMI-1640 supplemented with 20% fetal bovine serum (Sigma Chemical Co.) and 56.25 µg/mL phytohemagglutinin (Murex Diagnostics, Norcross, GA), and then stimulated by incubation at 37 °C with 5% CO2 for 72 hours. Only stimulated lymphocytes take up plasmids during transfection (21) and exhibit active nucleotide excision repair activity (23,24), which removes UV light-induced photoproducts from plasmids that harbor the reporter gene.
After 72 hours of stimulation, the cells from each subject were divided into four aliquots of approximately 1 x 106 cells each: two aliquots were transfected separately with untreated plasmids (as the baseline for comparison) and two aliquots were transfected separately with UV-irradiated plasmids. We used the diethylaminoethyl-dextran (Amersham Pharmacia Biotech, Inc., Piscataway, NJ) method (25) to transfect cells with approximately 0.25 µg of either untreated plasmid or plasmid that was pretreated with UV light (an incident UVC dose of 800 J/m2 at a wavelength of 254 nm from a 15 W unfiltered germicidal UVC lamp [Sankyo Denki Co. Ltd., Tokyo, Japan]). Cells were incubated at 37 °C with 5% CO2 after transfection.
We estimated CAT expression by measuring CAT enzyme activity in the transfected cells 40 hours after transfection as described previously (21). Briefly, the transfected cells were centrifuged at approximately 1200g for 10 minutes at room temperature; the cell pellets were collected and washed twice with 1.5 mL of Tris-buffered saline (TBS) and resuspended in 31.5 µL of 0.25 M TBS. The cells were lysed by three 10-minute cycles of freezing and thawing in a dry iceethanol bath and a 37 °C water bath. Cell extracts were then assayed for CAT expression or activity. The activity of the repaired CAT gene was determined by using a scintillation counter to measure [3H]monoacetylated and [3H]diacetylated chloramphenicols, which are formed by a reaction between chloramphenicol and [3H]acetyl coenzyme A that is catalyzed by the CAT protein in the cell extracts. DRC was defined as the ratio of the CAT activity of cells transfected with UV-treated plasmids (CATUV800) to that of cells transfected with untreated plasmids (CATUV0) multiplied by 100% (i.e., DRC = [CATUV800/CATUV0] x 100%). The CAT activity of cells transfected with undamaged plasmids provides an internal experimental control because it is derived under the same experimental conditions as the CATUV800 (24) and from the same number of cells from the same individual (21). The damage to the plasmids induced by UV before transfection was so substantial that some cells did not repair all plasmids after transfection, and thus DRC for cells transfected with damaged plasmids was always less than 100%. The actual residual CAT activity by cells bearing the damaged plasmids in this study was between 1% and 40% that of cells bearing undamaged plasmids. The coefficient of variation of repeated assays over 6 weeks is approximately 15% (21). The blastogenic rate (i.e., the percentage of lymphocytes that responded to phytohemagglutinin stimulation), cpm of CAT expression from the untreated plasmids (the baseline CAT expression level), and cell storage time (in months) were also recorded for comparisons between the case patients and the control subjects.
Statistical Analysis
The differences between case patients and control subjects in the distribution of demographic variables and known risk factors were examined by using the chi-square test. DRC was first analyzed as a continuous variable before and after natural logarithmic transformation to justify the need for data transformation. Students t test was used to compare differences in DRC between groups. Whenever the variance of the groups was statistically significant, Students t tests with unequal variances were used for comparisons. Correlation analyses were performed for DRC and selected variables. We used the median DRC of control subjects (9.40%) as the cutoff value to calculate crude odds ratios (ORs) and 95% confidence intervals (CIs). Values greater than the median value (9.40%) were considered to reflect proficient repair capacity, and values less than or equal to the median value (9.40%) were considered to reflect deficient (i.e., suboptimal) repair capacity. In the questionnaire, skin color was self-assessed on a scale of 1 (light) to 10 (dark); skin color values of 1 or 2 were defined as fair skin, values of 3 or 4 were defined as brown skin, and values of 510 were defined as dark skin.
Data on the Clark level for each tumor, which uses a scale of IV (where higher numbers indicate a deeper and more invasive melanoma) (26), were obtained from patients medical charts and used to examine whether DRC was associated with tumor phenotype. We assumed that tumors that occurred on sun-exposed skin were induced by sunlight and that tumors on unexposed skin were not. Sun-exposed skin included both habitually sun-exposed sites (such as the neck, arms, face, and skull) and intermittently sun-exposed sites (such as the feet, legs, upper back, and shoulders); unexposed skin included the chest/mantle, groin, pelvis, hip, or trunk (including abdomen). For logistic regression analysis, dummy variables of risk factor classes and the median and tertile values for DRC were created to calculate the ORs and 95% CIs. Adjusted ORs were calculated by fitting unconditional logistic regression models with adjustment for age, sex, and covariates such as known risk factors for CMM and assay-related variables (i.e., blastogenic rate, cell storage time, and baseline level of CAT activity). To perform the linear trend test, the dummy variables were recoded as continuous variables with values of 13 and fitted into an unconditional logistic regression model with and without adjustment for covariates. All statistical analyses were performed with the use of Statistical Analysis System software (version 8.0; SAS Institute, Inc., Cary, NC). All statistical tests were two-sided.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
As shown in Table 1, there were no statistically significant differences in the frequency distributions of age (chi-square test: P = .332) or sex (chi-square test: P = .634) between case patients and control subjects, suggesting adequate matching. There were also no statistically significant differences in marital status, educational level, or household income between case patients and control subjects (Table 1
), nor were there differences in smoking status (P = .500), use of hormone replacement therapy (in women) (P = .516), vitamin supplementation (P = .247), or history of exposure to medical radiation (P = .941) (data not shown). Although case patients reported using sunscreen more frequently than did control subjects (P = .001), this variable was not associated with DRC (data not shown). Among the case patients, 61% reported a family history of any cancer and 12% reported a family history of melanoma, whereas among control subjects, 56% reported a family history of any cancer and 9.3% reported a family history of melanoma. Neither of these differences was statistically significant (P = .237 for any cancer and P = .385 for melanoma) (data not shown).
|
|
|
|
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Although numerous epidemiologic studies have established that sunlight, fair complexion, moles or multiple atypical nevi, and family history of cancer are risk factors for CMM, few studies have investigated DRC as an etiologic factor for CMM in humans. Perhaps the best supporting evidence for the roles of DRC and sunlight in the etiology of CMM comes from XP patients who have germline mutations in genes involved in nucleotide excision repair, which is responsible for removing UV-induced DNA damage (27). In a study of 132 XP patients, 22% had CMM, and the risk of CMM in XP patients younger than 20 years was increased more than 1000-fold compared with the general population (20).
That CMM occurs frequently in the presence of XP or defective DRC is clear biologic evidence that low DRC plays a role in the etiology of CMM, but such a role in the general population is still controversial. However, several lines of evidence support this role of DRC in the general population. UV light induces melanocytic proliferation in both exposed and shielded areas of the skin (28), suggesting that melanocytes are responsive to UV exposure. Results of early laboratory studies on fibroblasts and lymphoblastoid cell lines derived from patients with and without CMM or dysplastic nevi, a precursor of CMM, suggested that patients with familial CMM are abnormally sensitive to UV light (29). Later studies revealed that this UV sensitivity may be due to a defect in the repair of UV-induced DNA damage (30). DNA mutations that are induced by UV (typically CCTT or C
T) have also been found (although relatively infrequently) in p16/MTS1/CDKN2A (31), a gene that is frequently mutated in familial melanomas (32). Furthermore, laboratory and animal studies have shown that unrepaired UV-induced photoproducts initiate systemic immunosuppression that contributes to cutaneous tumorigenesis (33), which may explain the occurrence, observed in this study, of CMM on unexposed skin.
Several studies have found no association between DRC and CMM. In a pilot casecontrol study, Hansson and Loow (25) found no statistically significant difference in the mean DRCs between lymphocytes from 17 patients with hereditary dysplastic nevi and lymphocytes from 17 healthy control subjects. Recently, in a larger study of 132 CMM case patients and 145 control subjects, it was also found that DRC was not an independent risk factor for CMM in an Italian population that had a high prevalence of dysplastic nevi (13). This study was previously the largest published study of DRC in CMM (13). There are several possible reasons for the discrepancy between the results of these two studies and our results. The majority of our case patients are thought to have had sporadic CMM, which should have a stronger environmental etiologic component than familial CMM. Low DRC is most likely to play a role in the etiology of sunlight-related CMM. Our study included more than twice as many subjects than the Italian study. Compared with the Italian study, we observed a larger variation in DRC among the control subjects (which may reflect either an inherent characteristic of the assay or true interindividual variation) and a smaller variation in DRC among the case patients (which may reflect selection of these subjects by risk factors such as sun exposure). However, our larger sample size substantially increased the power of our study. The discrepancy between the results of the Italian study and our study may also be due to lower sunlight exposure among the Italian population than among our Texas study population, because low DRC is not associated with CMM in the absence of sunlight exposure, even in XP patients. Tanning ability and the presence of dysplastic nevi, coupled with low DRC, were associated with an elevated risk of CMM in the Italian population, suggesting that DNA repair may indeed have a role in the etiology of CMM in these subgroups (13). There also may have been differences in the populations age distributions and lifetime sunlight exposure, the UV dose used to damage the plasmids, and the genetic backgrounds of the two study populations.
It has been suggested that sunlight exposure may cause nonmelanoma skin cancer and CMM in XP patients by different mechanisms (20), because there are differences in risks of CMM (which arises in melanocytes) and nonmelanoma skin cancer (which arises in keratinocytes) in response to UV in terms of repair rate or the ability of cells to survive with DNA damage. Whereas keratinocytes are prone to UV-induced apoptosis, melanocytes are more resistant because they express melanin, which protects against UV exposure, and because they constitutively express the anti-apoptotic protein Bcl2 (34). Therefore, although melanocytes are more resistant to the lethal effects of UV light, they are more likely to survive with mutations resulting from unrepaired DNA damage (35). Thus, DRC is likely to play an important role in UV-induced carcinogenesis in melanocytes.
Our study has some inherent limitations. First, because it was a hospital-based casecontrol study, the reported information on known risk factors, particularly family history of cancer, may not be representative of the general population. Second, we did not perform skin examinations of the control subjects and therefore we could not assess how they differed from case patients with regard to clinical skin type. Third, the DRC of lymphocytes (the cells in which our measurements were made) may differ from the DRC of melanocytes. Fourth, we cannot exclude the possibility that the disease state influences DRC. However, studies of XP patients provide supporting evidence that impaired DRC is associated with the development of skin cancers (20) and that low DRC, as a genetic trait, can be measured in any type of tissue (21). Our data also suggest that our assay results are valid in a biologic sense and were not affected by tumor or assay conditions. For instance, there was no apparent relationship between DRC and tumor thickness and level of invasion, indicating that DRC measured in lymphocytes was not affected by tumor status. Because DRC was measured by the host-cell reactivation assay using frozen cells assayed in batches, the effects of freezing and storage of the cells on DRC should be reflected in the blastogenic responses after mitogen stimulation. However, these assay correlates were not different between case patients and control subjects. Although the case samples were stored longer than the control samples, the cell storage time did not appear to have a substantial effect on lymphocyte function and DRC.
The results of this study are consistent with those of our previous study in which we found that low DRC (measured by the same host-cell reactivation assay) in peripheral lymphocytes is a risk factor for basal cell carcinoma (36). In the current study, we did not observe a previously described age-related decline in DRC (36), possibly because of differences in the age range of the subjects included. More mechanistic studies are needed to unravel the molecular mechanisms underlying the sex difference in DRC observed consistently in this and other studies (36,37).
![]() |
NOTES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We thank Dr. Larry Grossman (The Johns Hopkins University) for providing pCMVcat and scientific advice, Linda Young and Margaret Lung for subject recruitment, Dr. Lie Cheng and Yongli Guan for technical assistance, and Yolanda Bell and Joanne Sider for assistance in preparing the manuscript.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1 American Cancer Society, Inc. Cancer facts & figures 2002. Atlanta (GA): American Cancer Society; 2002. p. 4.
2 Glass A, Hoover RN. The emerging epidemic of melanoma and squamous cell skin cancer. JAMA 1989;262:2097100.[Abstract]
3 Franceschi S, La Vecchia C, Negri E, Levi F. Increases in mortality from cutaneous malignant melanoma in southern Europe. Int J Cancer 1992;51:1602.[Medline]
4 MacLennan R, Green AC, McLeod GR, Martin NG. Increasing incidence of cutaneous melanoma in Queensland, Australia. J Natl Cancer Inst 1992;84:142732.[Abstract]
5 Moan J, Dahlback A. The relationship between skin cancer, solar radiation and ozone depletion. Br J Cancer 1992;65:91621.[Medline]
6 Wingo PA, Ries LA, Rosenberg HM, Miller D, Edwards BK. Cancer incidence and mortality, 19731995. A report card for the U.S. Cancer 1998;82:1197207.[CrossRef][Medline]
7 Fears TR, Bird CC, Guerry D 4th, Sagebiel RW, Gail MH, Elder DE, et al. Average midrange ultraviolet radiation flux and time outdoors predict melanoma risk. Cancer Res 2002;62:39926.
8 Zanetti R, Franceschi S, Rosso S, Colonna S, Bidoli E. Cutaneous melanoma and sunburns in childhood in a Southern European population. Eur J Cancer 1992;28A:11726.[CrossRef]
9 Azizi E, Wax Y, Lusky A, Kushelevsky A, Schewach-Millet M. The recovery from ultraviolet radiation-induced erythema and melanoma risk factors: a case-control study. J Am Acad Dermatol 1990;23:306.[Medline]
10 Khlat M, Vail A, Parkin M, Green A. Mortality from melanoma in migrants to Australia: variation by age at arrival and duration of stay. Am J Epidemiol 1992;135:110313.[Abstract]
11 Green A, Sorahan T, Pope D, Siskind V, Hansen M, Hanson L, et al. Moles in Australian and British schoolchildren. Lancet 1988;2:1497.
12 Tucker MA, Halpern A, Holly EA, Hartge P, Elder DE, Sagebiel RW, et al. Clinically recognized dysplastic nevi. A central risk factor for cutaneous melanoma. JAMA 1997;277:143944.[Abstract]
13 Landi MT, Baccarelli A, Tarone RE, Pesatori A, Tucker MA, Hedayati M, et al. DNA repair, dysplastic nevi, and sunlight sensitivity in the development of cutaneous malignant melanoma. J Natl Cancer Inst 2002;94:94101.
14 Gallagher RP, McLean DI, Yang CP, Coldman AJ, Silver HK, Spinelli JJ, et al. Anatomic distribution of acquired melanocytic nevi in white children. A comparison with melanoma: The Vancouver Mole Study. Arch Dermatol 1990;126:46671.[Abstract]
15 Halpern AC, Cuerry D IV, Elder DE, Clark WH Jr, Synnestvedt M, Norman S, et al. Dysplastic nevi as risk markers of sporadic (nonfamilial) melanoma. Arch Dermatol 1991;127:9959.[Abstract]
16 Greene MH, Clark WH, Tucker MA, Elder DE, Kraemer KH, Guerry D 4th, et al. Acquired precursor of cutaneous malignant melanoma: the familial dysplastic nevus syndrome. N Engl J Med 1985;312:917.[Medline]
17 Tucker MA, Fraser MC, Goldstein AM, Elder DE, Guerry D 4th, Organic SM. Risk of melanoma and other cancers in melanoma-prone families. J Invest Dermatol 1993;100:350S355S.[Abstract]
18 Piepkorn M. Melanoma genetics: an update with focus on the CDKN2A(p16)/ARF tumor suppressors. J Am Acad Dermatol. 2000;42:70526.[Medline]
19 Platz A, Ringborg U, Hansson J. Hereditary cutaneous melanoma. Semin Cancer Biol 2000;10:31926.[CrossRef][Medline]
20 Kraemer KH, Lee MM, Andrews AD, Lambert WC. The role of sunlight and DNA repair in melanoma and nonmelanoma skin cancer. The xeroderma pigmentosum paradigm. Arch Dermatol 1994;130:101821.[Abstract]
21 Athas AF, Hedayati M, Matanoski GM, Farmer ER, Grossman L. Development and field-test validation of an assay for DNA repair in circulating human lymphocytes. Cancer Res 1991;51:578693.[Abstract]
22 Protic-Sabljic M, Kraemer KH. One pyrimidine dimer inactivates expression of a transfected gene in xeroderma pigmentosum cells. Proc Natl Acad Sci U S A 1985;82:66226.[Abstract]
23 Shivji MK, Kenny MK, Wood R. Proliferating cell nuclear antigen is required for DNA excision repair. Cell 1992;69:36774.[Medline]
24 Barret JM, Calsou P, Salles B. Deficient nucleotide excision repair activity in protein extracts from normal human lymphocytes. Carcinogenesis 1995;16:16116.[Abstract]
25 Hansson J, Loow H. Normal reactivation of plasmid DNA inactivation by UV irradiation by lymphocytes from individuals with hereditary dysplastic naevus syndrome. Melanoma Res 1994;4:1637.[Medline]
26 Busam KJ. Lack of relevant information for tumor staging in pathology reports of primary cutaneous melanoma. Am J Clin Pathol 2001;115:7436.[Medline]
27 Cleaver JE, Crowley E. UV damage, DNA repair and skin carcinogenesis. Front Biosci 2002;7:d102443.[Medline]
28 Stierner U, Rosdahl I, Augustsson A, Kagedal B. UVB irradiation induced melanocyte increase in both exposed and shielded human skin.J Invest Dermatol 1989;92:5614.[Abstract]
29 Roser M, Bohm A, Oldigs M. Ultraviolet-induced formation of micronuclei and sister chromatid exchange in cultured fibroblasts of patients with cutaneous malignant melanoma. Cancer Genet Cytogenet 1989;41:12937.[Medline]
30 Moriwasi SI, Tarone RE, Tucker MA, Goldstein AM, Kraemer KH. Hypermutability of UV-treated plasmids in dysplastic nevus/familial melanoma cell lines. Cancer Res 1997;57:463741.[Abstract]
31 Kumar R, Rozell BL, Louhelainen J, Hemminki K. Mutations in the CDKN2A(p16INK4a) gene in microdissected sporadic primary melanomas. Int J Cancer 1998;75:1938.[CrossRef][Medline]
32 Hussussian CJ, Struewing JP, Goldstein AM, Higgins PA, Ally DS, Sheahan MD, et al. Germline p16 mutations in familial melanoma. Nat Genet 1994;8:1521.[Medline]
33 Kripke ML, Cox PA, Alas LG, Yarosh DB. Pyrimidine dimers in DNA initiate systemic immunosuppression in UV irradiated mice. Proc Natl Acad Sci U S A 1992;89:751620.[Abstract]
34 Plettenberg A, Ballaun C, Pammer J, Mildner M, Strunk D, Weninger W, et al. Human melanocytes and melanoma cells constitutively express the Bcl-2 proto-oncogene in situ and in cell culture. Am J Pathol 1995;146:6519.[Abstract]
35 De Leeuw SM, Janssen S, Simons JW, Lohman PH, Vermeer B. UV action spectrum for the clone-forming ability of cultured human melanocytes and keratinocytes. Photochem Photobiol 1994;59:4306.[Medline]
36 Wei Q, Matanoski GM, Farmer ER, Hedayati MA, Grossman L. DNA repair and aging in basal cell carcinoma: a molecular epidemiology study. Proc Natl Acad Sci U S A 1993;90:16148.[Abstract]
37 Wei Q, Cheng L, Amos CI, Wang LE, Guo ZZ, Hong WK, et al. Repair of tobacco carcinogen-induced DNA adducts and lung cancer risk: a molecular epidemiologic study. J Natl Cancer Inst 2000:92:176472.
Manuscript received June 7, 2002; revised November 26, 2002; accepted December 9, 2002.
This article has been cited by other articles in HighWire Press-hosted journals:
![]() |
||||
|
Oxford University Press Privacy Policy and Legal Statement |