The nucleotide metabolism enzyme ribonucleotide reductase is composed of a regulatory subunit (RRM1) and a catalytic subunit (RRM2). The RRM1 locus has frequent loss of heterozygosity in lung cancers, ectopic expression of RRM1 suppresses proliferation of ras-transformed mouse fibroblasts, and high levels of RRM1 expression are associated with a significant survival benefit in patients with lung cancer. In RRM1 transgenic human lung and colon cancer cell lines, we observed induction of G2 cell cycle arrest, apoptosis, and efficient DNA damage repair. We generated strains of RRM1 transgenic mice and found that carcinogen-induced lung tumor formation was significantly suppressed. The tumor suppression was more pronounced in strains with high levels of RRM1 expression than in those with low levels of expression. DNA damage repair capacity in transgenic animals was determined, and RRM1 transgenic animals repaired chemically induced DNA damage with greater efficiency than control animals. We conclude that the regulatory subunit of ribonucleotide reductase has tumor suppressor activity that is mediated through efficient DNA damage repair. (Cancer Res 2006; 66(13): 6497-502)

Ribonucleotide reductase is required for DNA synthesis and repair in mammalian cells. It catalyzes the rate-limiting step in deoxynucleotide formation and is composed of two regulatory subunits (RRM1) and two catalytic subunits (RRM2 or p53R2). The RRM1 subunit controls substrate specificity and on/off function of ribonucleotide reductase, whereas the catalytic subunit converts nucleotides to deoxynucleotides (14). The catalytic subunit (RRM2) has a half-life (t1/2) of 3 hours, and measurable levels are detectable during late G1 and S phase. The regulatory subunit (RRM1) is present throughout the cell cycle in relatively constant amounts and its t1/2 is ∼15 hours (5, 6), which suggests that RRM1 may have cellular functions beyond its role in ribonucleotide reductase formation.

Three independent lines of evidence suggest a role for RRM1 as a tumor suppressor. First, the RRM1 locus on human chromosome 11p15.5 has frequent loss of heterozygosity in patients with lung cancer (7), and high levels of RRM1 expression are significantly correlated with long survival of patients (8). Second, stable expression of RRM1 in a ras-transfected mouse fibroblast cell line resulted in reduced anchorage-independent growth and tumor formation in syngeneic mice (9). Third, induction of RRM1 expression in human lung cancer cell lines at physiologic levels resulted in decreased cellular migration and invasion; in animal models, increased RRM1 expression in syngeneic lung cancers suppressed metastasis formation and increased survival (10).

These observations led us to investigate if RRM1 would trigger cellular responses associated with DNA damage checkpoint induction and if it would suppress carcinogen-induced tumor formation in a transgenic animal model.

Cell lines and constructs. We cloned the complete open reading frame of 2,379 nucleotides of the human RRM1 gene into the SrfI and XhoI sites of the mammalian expression vector pCMV-Tag2 in frame (pCMV-R1) and for control purposes out-of-frame (pCMV-Ct). Both constructs were verified by sequencing in both directions. These plasmids were transfected into the human lung and colon cancer cell lines NCI-H23 and HCT-8, respectively, using cationic liposomes. Stable transfectants were isolated by G418 selection. Clonal sublines were generated by serial dilution in 96-well cluster plates. Southern blot analysis with a human RRM1 probe of HindIII- and BamHI-digested genomic DNA from these cell lines was used to confirm stable transgene integration into the genome. An additional 2.4-kb band representing the RRM1 transgene was present in the transfected cell lines, whereas it was absent in the parent cell lines.

Real-time quantitative reverse transcription-PCR and immunoblotting. Real-time quantitative reverse transcription-PCR (RT-PCR) was used to quantify target gene expression. For each of the target genes, we designed custom primers and probes. All primer pairs were intron spanning to avoid amplification of potentially contaminating genomic DNA, and all were verified by bidirectional sequencing of amplified cDNA products. Commercially available primers and probes were used to quantify the housekeeping genes 18SrRNA and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as internal references. They have been selected among 10 genes as a result of a screen for the best reference genes through determination of differential expression at various concentrations of RNA. The equivalent of 5 ng total RNA was used as template for each RT-PCR. For standard curve determination, a mixture of cDNAs derived from a random selection of cell lines was used. Amplifications were done in 96-well plates, and each sample was run in triplicate. Each plate included samples for a standard curve and negative controls without template cDNA. Fluorescent emission resulting from probe cleavage was recorded in real-time on an ABI Prism 7700 Sequence Detection System (Perkin-Elmer, Foster City, CA).

Immunoblot analysis was done on whole-cell lysates prepared from exponentially growing cultured cells or freshly frozen tissue. Protein content was determined with the Bio-Rad detergent-compatible protein assay (Bio-Rad, Hercules, CA). Protein (100 μg) was loaded per lane, separated in SDS polyacrylamide gels, and transferred to membranes. Target proteins were labeled using specific, commercially available antisera or antibodies [RRM1 (Cell Signaling Technology, Inc., Danvers, MA), GADD45 (Chemicon International, Inc., Temecula, CA), AKT, pAKT(473), pAKT(308), poly(ADP-ribose) polymerase (PARP; 116; Cell Signaling Technology), and β-actin (Sigma-Aldrich, Inc., St. Louis, MO)], and they were visualized by chemiluminescence and autoradiography. Signal intensities for the respective bands were measured by densitometry and normalized to the β-actin signal.

Time-lapse videography. Time-lapse videography (TLV) was done in T25 flasks with the video camera in the incubator and focused on an area that contained ∼10 to 15 cells. A temperature probe was used to ensure that the culture medium did not exceed 38°C as a result of the videography. Continual recording was done for 10 days. Mitoses of each cell were observed, and the time elapsed between two events was recorded.

Cell cycle distribution analysis. Fluorescence activated cell sorting of ∼100,000 exponentially growing cells stained with propidium iodine was used to determine the distribution of cells in separate phases of the cell cycle. To distinguish between G2 and M phase, mithramycin staining was used (11). The percentage of cells in G2-M phase was compared with the expression of RRM1 in clonal lines derived from H23-R1.

Spectral karyotyping. Well-separated mitotic chromosomes were prepared by standard techniques. They were hybridized to paint probes consisting of chromosome-specific sequences labeled with varying amounts of dyes such that each chromosome pair has unique spectral characteristics (Vysis, Abbott Molecular, Inc., North Chicago, IL). Color variations were detected by a computer program that reassigns visually distinct colors to each chromosome pair. Slides were then mounted with 4′,6-diamidino-2-phenylindole in antifade solution. Fifty metaphases were analyzed from H23-R1, H23-Ct, and NCI-H23 for numerical and structural chromosomal aberrations.

Comet assay. Single-cell suspensions were prepared and exposed for 10 minutes to 0.1 mmol/L H2O2 in six-well cluster plates containing 100,000 cells per well. They were washed and embedded in 0.5% low-melt agarose on glass slides. The embedded cells were lysed (2.5 mol/l NaCl, 100 mmol/L EDTA, 10 mmol/L Tris-base, 1% sodium lauryl sarcosine, 0.01% Triton X-100), 30 minutes for cells and 60 minutes for splenocytes, and denatured (0.3 mol/l NaOH, 1 mmol/L EDTA). The slides were then transferred to Tris-borate EDTA buffer (90 mmol/L Tris-borate, 2 mmol/L EDTA). An electric field, 1 V/cm, was applied for 15 minutes similar to a standard agarose gel electrophoresis. In this assay, damaged DNA will “leak” out of the nucleus and form a “comet” in the direction of the electric field, whereas intact DNA will remain nuclear (12). The size and number of comets is proportional to the amount of DNA damage, and it can be measured using an image analysis software program (Comet Analysis software, Loats Associates, Westminster, MD). The assay allows for a global assessment of DNA damage repair capacity. For this, cells were allowed to repair chemical- or radiation-induced DNA damage for various periods before embedding in agarose and electrophoresis.

Generation of RRM1 transgenic mouse strains. Pronuclei of FVB/n mice were injected with a DNA fragment that contained the complete human RRM1 gene under the control of a cytomegalovirus promoter. Human RRM1 has 90.3% identity with the mouse gene at the nucleotide level and 97.7% identity at the amino acid level. Only 18 amino acids are different between these species (Genbank accession nos. are X59617 for the human gene and NM_009103 for the mouse gene). Three founder animals were identified that produced offspring by crossbreeding with wild-type FVB/n mice (isogenic strains 35, 167, and 168). Incorporation of the human RRM1 gene into the mouse genome was verified by analysis of genomic DNA for transgene-specific restriction fragments [transgenic-positive (tg+) mice].

Carcinogen-induced lung tumor formation and animal survival. The carcinogenesis model used was the urethane model, where a simple carcinogen induces cancer after application of a single dose (13). The model is well established and has been used to investigate lung carcinogenesis. Approximately equal numbers of tg+ and transgenic-negative (tg−) animals received a single i.p. injection of urethane at a dose of 1 mg/g body weight in 0.9% NaCl at the age of 6 to 8 weeks. For evaluation of the effect of RRM1 transgene expression on lung tumor formation, animals were euthanized at the age of 6 months by CO inhalation, a method recommended by the American Veterinary Medical Association panel on euthanasia. Organs were harvested and immediately frozen in liquid nitrogen or fixed in formaldehyde. Standard H&E-stained sections of lung were viewed, and the number of pulmonary adenomas and carcinomas was counted. For evaluation of the effect of RRM1 transgene expression on natural animal survival, equal numbers of tg+ and tg− animals were allowed to age and die of natural causes. The time elapsed from birth to death was recorded in these animals.

Statistical analysis. Spearman's rank-order calculation was used to test for an association between the level of RRM1 expression and the percentage of cells in G2 phase of the cell cycle. To test for a difference in the number of tumors between tg+ and tg− animals, the χ2 test was used. The number of tumors per animal was assessed in a blinded fashion. To analyze animal survival, the time from birth to natural death was recorded for each animal. Those that had not died at the time of conclusion of the experiment were censored as of that date. Kaplan-Meier survival curves were generated for tg+ and tg− animals, and the difference in survival was tested for statistical significance with the log-rank test.

We generated stable human RRM1 transfectants of the human lung cancer cell line NCI-H23 (H23-R1) and the colon cancer cell line HCT-8 (HCT8-R1). As a control, we transfected the same cell lines with a plasmid that contained an out-of-frame RRM1 construct (H23-Ct and HCT8-Ct; ref. 10).

TLV was done to continually record cellular divisions of 10 to 15 cells and their progeny over 10 days. The mean interdivision time for H23-R1 was 41.9 hours [SD, 19 hours; 95% confidence interval (95% CI), 32.3-51.5 hours], and it was 25.7 hours for H23-Ct (SD, 3.9 hours; 95% CI, 24.3-27.1 hours; Fig. 1; the video can be viewed at http://researchdata.moffitt.usf.edu/thoracic/RRM1-H23-TLV-0006.wmv).

Figure 1.

Schema of interdivision times by TLV. A, H23-Ct (relative RRM1 expression 1). The progeny of 14 individual cells were followed. X, two cells died. Cells 2, 4, and 9 moved out of the view area. The example shown is from cell 8, which is representative of all nine others that produced clones. The interdivision time ranged from 12.6 to 37.6 hours (mean, 25.7 hours; SD, 3.9 hours; 95% CI, 24.3-27.1 hours). B, H23-R1 (relative RRM1 expression 2.2). The progeny of 13 individual cells were followed. Eight cells died (X). Five cells (1, 3, 6, 7, and 12) successfully divided. The interdivision time among these five clones ranged from 8.3 to 89.3 hours (mean, 41.9 hours; SD, 19 hours; 95% CI, 32.3-51.5 hours).

Figure 1.

Schema of interdivision times by TLV. A, H23-Ct (relative RRM1 expression 1). The progeny of 14 individual cells were followed. X, two cells died. Cells 2, 4, and 9 moved out of the view area. The example shown is from cell 8, which is representative of all nine others that produced clones. The interdivision time ranged from 12.6 to 37.6 hours (mean, 25.7 hours; SD, 3.9 hours; 95% CI, 24.3-27.1 hours). B, H23-R1 (relative RRM1 expression 2.2). The progeny of 13 individual cells were followed. Eight cells died (X). Five cells (1, 3, 6, 7, and 12) successfully divided. The interdivision time among these five clones ranged from 8.3 to 89.3 hours (mean, 41.9 hours; SD, 19 hours; 95% CI, 32.3-51.5 hours).

Close modal

We did a cell cycle distribution analysis. The proportion of cells in G2 phase was significantly higher in H23-R1 (27%) and HCT8-R1 (17.7%) compared with H23-Ct (6%) and HCT8-Ct (1.9%; Fig. 2). That cells were in G2 and not in M phase was confirmed by counting metaphases manually and by fluorescence-activated cell sorting (FACS) analysis after mithramycin staining (11). There was no difference in the number of cells in M phase between these lines, suggesting the P-value was that RRM1 triggers a delay in G2 progression. Clonal sublines of H23-R1 were generated by serial dilution, which expressed RRM1 at various levels. There was a significant correlation between the percentage of cells in G2 and the relative expression of RRM1 in these sublines as determined by quantitative real-time RT-PCR (r = 0.688; P = 0.013, Spearman's rank; Fig. 2C).

Figure 2.

Cell cycle distribution using propidium iodine staining and FACS analysis. A, H23-Ct (relative RRM1 expression 1). Percentage of cells in G1 (42%), S (51.6%), and G2-M (6.4%). B, H23-R1 (relative RRM1 expression 2.2). Percentage of cells in G1 (16.6%), S (56.6%), G2-M (26.8%). C, correlation between the relative RRM1 expression in H23-Ct, H23-R1, and 10 subclones of H23-R1 and the proportion of cells in G2 phase of the cell cycle. Spearman's rank correlation coefficient was 0.688, and the P-value was 0.013.

Figure 2.

Cell cycle distribution using propidium iodine staining and FACS analysis. A, H23-Ct (relative RRM1 expression 1). Percentage of cells in G1 (42%), S (51.6%), and G2-M (6.4%). B, H23-R1 (relative RRM1 expression 2.2). Percentage of cells in G1 (16.6%), S (56.6%), G2-M (26.8%). C, correlation between the relative RRM1 expression in H23-Ct, H23-R1, and 10 subclones of H23-R1 and the proportion of cells in G2 phase of the cell cycle. Spearman's rank correlation coefficient was 0.688, and the P-value was 0.013.

Close modal

We used spectral karyotyping to assess if differential numerical or structural chromosome aberrations are present in NCI-H23, H23-R1, and H23-Ct. No novel or unique aberrations were observed in H23-R1, and all three lines were indistinguishable by spectral karyotyping. However, there was a significantly higher number of aberrations with this technique than expected from previous G-banding analyses. The only chromosome not involved in any translocation was chromosome 12. All three cell lines contained two mutually exclusive clonal abnormalities, t(15:X) and t(17:X).

Because GADD45 is a key gene involved in G2 checkpoint induction (14, 15), immunoblotting and quantitative real-time RT-PCR was used to assess its expression. H23-R1 and HCT8-R1 cells expressed GADD45 at higher levels than H23-Ct and HCT8-Ct cells (Fig. 3), suggesting that the observed RRM1-induced G2 cell cycle arrest is mediated by GADD45.

Figure 3.

Immunoblot for expression of GADD45, AKT, pAKT, and PARP in relation to RRM1 expression in transfected cell lines. H23, parent cell line; H23-Ct, control transfected H23; and H23-R1, RRM1-transfected H23. The relative expression in H23-R1 compared with H23-Ct corrected for actin was 1.54-fold for RRM1, 3.85-fold for GADD45, 0.33- to 0.45-fold for pAKT, and 0.05-fold for PARP.

Figure 3.

Immunoblot for expression of GADD45, AKT, pAKT, and PARP in relation to RRM1 expression in transfected cell lines. H23, parent cell line; H23-Ct, control transfected H23; and H23-R1, RRM1-transfected H23. The relative expression in H23-R1 compared with H23-Ct corrected for actin was 1.54-fold for RRM1, 3.85-fold for GADD45, 0.33- to 0.45-fold for pAKT, and 0.05-fold for PARP.

Close modal

We assessed if the G2 arrest would affect DNA damage repair efficiency. H23-R1 and H23-Ct cells were exposed to DNA-damaging agents here, H2O2 and ionizing radiation, and the amount of damage was assessed with the comet assay (12). The level of RRM1 expression did not affect the initial amount of DNA damage; however, it significantly affected the efficiency with which the damage was repaired (Fig. 4A). In the example shown, a complete reversal of measurable comets occurred after 2 hours of repair in H23-R1 cells. In contrast, in H23-Ct cells, complete comet reversal was only seen after 24 hours, suggesting that RRM1 not only delays G2 cell cycle progression but also increases DNA damage repair efficiency.

Figure 4.

DNA damage and repair in RRM1 transgenic and nontransgenic cells as determined by the comet assay. Cells were treated for 10 minutes with 0.1 mmol/L H2O2 in six-well cluster plates containing 100,000 cells. A, measurable DNA damage (comet moments) in H23-Ct and H23-R1 after H2O2 exposure and 2 and 24 hours after removal of the damaging agent. H23-R1 cells have completely reversed DNA damage after 2 hours, whereas H23-Ct cells require 24 hours for complete repair. B, measurable DNA damage in splenocytes from RRM1 tg+ and tg− strain 35 mice after H2O2 exposure and 2 to 60 hours after removal of the damaging agent. 35tg+ cells have repaired DNA damage after 24 hours, whereas 35tg− cells require >60 hours for repair. C, strain 35 tg− splenocyte after H2O2 exposure without repair. D, strain 35 tg− splenocyte after 24 hours of repair following H2O2 exposure. E, strain 35 tg+ splenocyte after H2O2 exposure without repair. F, strain 35 tg+ splenocyte after 24 hours of repair following H2O2 exposure.

Figure 4.

DNA damage and repair in RRM1 transgenic and nontransgenic cells as determined by the comet assay. Cells were treated for 10 minutes with 0.1 mmol/L H2O2 in six-well cluster plates containing 100,000 cells. A, measurable DNA damage (comet moments) in H23-Ct and H23-R1 after H2O2 exposure and 2 and 24 hours after removal of the damaging agent. H23-R1 cells have completely reversed DNA damage after 2 hours, whereas H23-Ct cells require 24 hours for complete repair. B, measurable DNA damage in splenocytes from RRM1 tg+ and tg− strain 35 mice after H2O2 exposure and 2 to 60 hours after removal of the damaging agent. 35tg+ cells have repaired DNA damage after 24 hours, whereas 35tg− cells require >60 hours for repair. C, strain 35 tg− splenocyte after H2O2 exposure without repair. D, strain 35 tg− splenocyte after 24 hours of repair following H2O2 exposure. E, strain 35 tg+ splenocyte after H2O2 exposure without repair. F, strain 35 tg+ splenocyte after 24 hours of repair following H2O2 exposure.

Close modal

The TLV also suggested increased apoptosis in H23-R1 cells. Using a DNA fragment end-labeling assay (terminal deoxynucleotidyl transferase–mediated dUTP nick end labeling; APOPercentage, Accurate Chemical and Scientific Corp., Westbury, NY), we confirmed increased apoptosis in H23-R1 compared with H23-Ct. In a series of immunoblotting analyses for proteins implicated in apoptosis, we found no significant differences between the parental cell line NCI-H23 and the control cell line H23-Ct. However, H23-R1 had a substantial increase for the proteins PARP p89 (2.2-fold; cleaved product of PARP p116), apoptosis-inducing factor (2.7-fold), and cytochrome c (1.7-fold) compared with H23-Ct. The total level of Akt was identical in both cell lines; however, Akt phosphorylation and uncleaved PARP p116 levels were substantially decreased in H23-R1 compared with H23-Ct (Fig. 3).

We reasoned that the RRM1-mediated increased DNA damage repair capacity might also be operational in normal epithelial cells and thus suppress carcinogen-induced malignant transformation. To test this hypothesis, we generated three RRM1 transgenic mouse strains (tg+ strains 35, 167, and 168) and studied carcinogen-induced lung tumor formation compared with isogenic, nontransgenic mice. RRM1 expression was measured in lung tissue collected from 26-week-old tg+ mice and tg− littermates that served as controls using quantitative real-time RT-PCR. The average RRM1 expression was >7-fold in tg+ strain 35, >4-fold in tg+ strain 167, and not altered in tg+ strain 168 compared with tg− control animals. The average expression for GAPDH, adjusted for 18SrRNA expression, was not different between tg+ and tg− animals. We did not observe gross phenotypic differences between transgenic and control mice.

A murine chemical carcinogenesis model was used. In this model, a single i.p. injection of urethane results in benign and malignant lung tumors, and it has been used to investigate lung carcinogenesis (13). Mice of all three strains were injected with a single dose (1 mg/g) of urethane i.p. at the age of 8 to 9 weeks, and a necropsy was done 16 to 19 weeks thereafter. All tissues were inspected for tumor formation, and selected organs were examined microscopically. The number of animals with lung tumors was significantly reduced in strain 35 (N = 68; P = 0.0025, χ2 test), whereas no reduction was noted in strain 168 (N = 88; P = 0.7315, χ2 test). Tumor reduction was intermediate in strain 167 (Table 1). We measured RRM1 expression in lung tumors collected from 26-week-old strain 35 animals and found a >5-fold expression in tg+ compared with tg− animals.

Table 1.

Lung Tumor Formation in RRM1 Transgenic and Control Animals

RRM1 expressionAnimals with tumor* (n)Animals without tumor (n)Animals with tumor (%)Animals without tumor (%)Change in tumor formation (%)
Strain 35 (N = 68)       
    RRM1 tg+ 6.8× 15 10 60 40 −34 
    RRM1 tg− Control 39 91  
Strain 167 (N = 22)       
    RRM1 tg+ 3.7× 60 40 −27 
    RRM1 tg− Control 14 82 18  
Strain 168§ (N = 88)       
    RRM1 tg+ 1× 28 13 68 32 
    RRM1 tg− Control 30 17 64 36  
RRM1 expressionAnimals with tumor* (n)Animals without tumor (n)Animals with tumor (%)Animals without tumor (%)Change in tumor formation (%)
Strain 35 (N = 68)       
    RRM1 tg+ 6.8× 15 10 60 40 −34 
    RRM1 tg− Control 39 91  
Strain 167 (N = 22)       
    RRM1 tg+ 3.7× 60 40 −27 
    RRM1 tg− Control 14 82 18  
Strain 168§ (N = 88)       
    RRM1 tg+ 1× 28 13 68 32 
    RRM1 tg− Control 30 17 64 36  
*

All lung tumors were bronchioloalveolar carcinomas or adenomas.

P = 0.0025, χ2 test.

P = 0.5482, Fisher's exact test.

§

P = 0.7315, χ2 test.

Ninety-nine strain 35 animals (49 tg+ and 50 tg−) were used to assess RRM1s effect on natural survival. Death occurred in 42 animals between the ages of 68 and 390 days. Fifty-seven animals were alive and between the ages of 263 and 418 days at the time of study termination. Kaplan-Meier survival curves were generated (Fig. 5). RRM1 tg+ mice had a significantly longer survival than RRM1 tg− animals (P = 0.015, log-rank test).

Figure 5.

Kaplan-Meier plots of RRM1 tg+ and tg− FVB/n mouse strain 35. Mice were injected with a single dose of the carcinogen urethane (1 mg/g body weight in 0.9% NaCl at an age of 58-64 days). Survival was measured as the number of days elapsed from birth to death. A total of 50 tg− and 49 tg+ mice were used. Of the tg− mice, 28 had died and 22 were censored; of the tg+ mice, 14 had died and 35 were censored. The median follow-up time for all animals was 287 days (261 days for tg− and 287 days for tg+). The median survival in tg− mice was 258 days, and it was >365 days for tg+ mice. The survival of tg+ mice was significantly better than the survival of tg− mice (P = 0.015, log-rank test).

Figure 5.

Kaplan-Meier plots of RRM1 tg+ and tg− FVB/n mouse strain 35. Mice were injected with a single dose of the carcinogen urethane (1 mg/g body weight in 0.9% NaCl at an age of 58-64 days). Survival was measured as the number of days elapsed from birth to death. A total of 50 tg− and 49 tg+ mice were used. Of the tg− mice, 28 had died and 22 were censored; of the tg+ mice, 14 had died and 35 were censored. The median follow-up time for all animals was 287 days (261 days for tg− and 287 days for tg+). The median survival in tg− mice was 258 days, and it was >365 days for tg+ mice. The survival of tg+ mice was significantly better than the survival of tg− mice (P = 0.015, log-rank test).

Close modal

We used splenocytes from strain 35 animals to determine if DNA damage repair was differentially affected in tg+ and tg− animals. Our results showed that tg+ animals repaired H2O2-induced damage within 24 hours of removal of the agent, whereas tg− animals required over 60 hours for damage repair (Fig. 4B-F).

Preliminary evidence had suggested that RRM1 may function as a tumor suppressor (710). We generated in vitro and in vivo models to corroborate this evidence and found that RRM1 can trigger a G2 cell cycle checkpoint response with increased DNA damage repair and apoptosis and suppression of carcinogenesis in transgenic animals.

Physiologically, cell cycle arrest in proliferating cells results from a checkpoint induction, which is triggered by DNA damage (16). In our in vitro model system, we found no evidence for increased constitutional DNA damage in the transgenic cell lines using spectral karyotyping and the comet assay. Our in vitro results show that forced expression of RRM1 in human epithelial malignancies, as shown for lung and colon cancer, engages the G2 checkpoint through induction of GADD45 expression with subsequent increase in the efficiency of DNA damage repair and induction of apoptosis. Our data suggest that the increase in RRM1-induced apoptosis is mediated through the endogenous mitochondrial pathway, which is a well-described cellular mechanism of extensive DNA damage (17).

DNA damage is thought to be a major cause of human cancer (18, 19). The damage can occur spontaneously and can be triggered by chemicals or ionizing radiation. In our transgenic animal model, we show that RRM1 is crucially involved in containing chemical-induced carcinogenesis through efficient DNA damage repair. RRM1 can suppress tumor formation as evidenced by the dose-dependent reduction in lung tumors. The biological significance of this is underlined by the demonstration that RRM1 transgenic animals have a significantly better life expectancy that control animals after carcinogen exposure. This observation is completely consistent with our previous report of a survival benefit in patients with lung cancer after surgical resection whose tumors express high levels of RRM1 (8).

One potential implication of our data for persons is that those with high constitutional RRM1 expression may be better protected against DNA damage induced by carcinogens than those with relatively low levels of expression. Another potential effect may be on therapeutic efficacy of DNA-damaging agents, such as radiation and chemotherapy, which are the present-day mainstay of cancer treatment. In fact, recent data have suggested a decrease in chemotherapeutic efficacy in patients with high tumoral expression of RRM1 (20). Prospective epidemiologic and clinical studies are needed to delineate RRM1s effect on carcinogenesis and therapy in humans.

Grant support: National Cancer Institute grant R01 CA102726.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Hartmut Berns, Alan Cantor, Robert Engelman, Noreen Luetteke, and Swati Sharma for their support in conducting this work.

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